Patent Application
20220218834
CD163 is a membrane receptor molecule expressed on macrophages and monocytes. In some cases, CD163 may be amongst the most highly expressed receptor on macrophages and monocytes and functions as an endocytic receptor for hemoglobin-haptoglobin complexes. In this role, CD163 is believed to take up more than 1 g of hemoglobin each day.
Various pathophysiological conditions can lead to states of hemolysis. Hemolysis is characterized by the rupture of red blood cells (RBCs), which releases toxic cell-free hemoglobin (Hb) into the circulation. In addition to cell-free Hb toxicity, the breakdown of cell-free Hb into heme and apoglobin leads to the toxic overload of free heme in vivo. Finally, breakdown of heme into iron, biliverdin and carbon monoxide can also lead to iron buildup in vivo. Hemolytic conditions include, but are not limited to, malaria, red blood cell transfusion, beta-thalassemia, sickle-cell anemia, severe burns, radiation poisoning, surgery, extracorporeal circulation and others. Currently, there lacks an effective treatment for states of hemolysis, with patients relying on blood-transfusions to replaced lost red blood cells without treatment of cell-free Hb, free heme and/or iron toxicity.
Macrophages and monocytes are part of the innate immune defence and play a central role in many infectious, autoimmune, and malignant diseases such as cancer. In autoimmune/inflammatory disease such as rheumatoid arthritis, macrophages and monocytes are the main source of inflammatory molecules such as TNF-alpha, known to be of crucial importance in disease progression. In many infectious diseases such as tuberculosis (TB) and HIV, macrophages and monocytes can also harbor infectious agents. A few malignant diseases have their origin in cells of the monocytic/macrophage lineage such as histiocytic sarcoma. Further macrophages have a central role in immune evasion of cancers and may be a key target for new and improved immunotherapies for cancer treatment.
Direct targeting of drugs to macrophages and monocytes (for example, to down-regulate production of inflammatory cytokines, to kill intracellular organisms, or to kill malignant cells) may therefore have significant impact on certain diseases without influencing other cells in the body. The targeting may therefore increase the therapeutic index of the drug.
Patients suffering from states of hemolysis may benefit from scavengers that can detoxify Hb, heme and/or free iron so that these toxic species can be neutralized and safely cleared from the body. Thus, development of new scavenger proteins for Hb, heme and iron is a promising treatment modality for states of hemolysis.
During pathophysiological conditions characterized by extensive hemolysis (e.g., sickle cell anemia, malaria, red blood cell transfusion, etc.), free heme and cell-free hemoglobin (Hb) are released into the blood stream. Once released, free heme and Hb cause a variety of side-effects such as vasoconstriction, hypertension, oxidative tissue injury and kidney damage. Thus, treatment of hemolytic conditions would benefit from scavengers of free heme and cell-free Hb such as hemopexin (Hx) and haptoglobin (Hp), respectively.
Apohemoglobin (apoHb) is a protein that is produced by removing heme from Hb. Therefore, the vacant heme-binding pockets of apoHb possess a high affinity for heme. While apoHb has shown heme-binding activity in vitro, its use for hemolysis treatment has not been explored.
A major issue with potential in vivo administration of apoHb as a heme scavenger would be its low thermal stability at physiological temperature and short circulatory half-life (would be equivalent to Hb dimers, on the order of 30 min). Similar to Hb, apoHb can react with Hp to form a stable protein complex. The apoHb-Hp complex is more stable at physiological temperature compared to free apoHb, and maintains its ability to bind heme. Further, the apoHb-Hp complex can not only scavenge heme via the bound apoHb, but could also scavenge free Hb by exchanging bound apoHb for Hb due to the irreversibility of Hb-Hp binding. These two potential mechanisms for treating the side products of hemolysis with this novel therapeutic are summarized in
ApoHb can also be used for drug delivery applications. Yet, these applications would rely on the presence of Hp in the plasma for targeted drug delivery of the apoHb-drug conjugate to macrophages and monocytes and could be deterred by the instability of free apoHb at physiological temperature. Binding the apoHb-drug conjugate to Hp could prevent these issues and improve drug delivery to CD163+ macrophages and monocytes. Targeting CD163+ macrophages and monocytes would be beneficial under conditions of inflammation which induce high expression of CD163 receptors on the surface of macrophages and monocytes. Additionally, certain types of cancers (such as breast cancer) have tumor associated macrophages and monocytes with high CD163 expression which could facilitate targeted drug delivery.
A benefit for therapeutics/diagnostics delivered via apoHb-Hp is the potential for long circulatory half-life. Although Hp-Hb complexes can be quickly removed from the circulation due to the high specificity of CD163+ macrophage capture, saturation of these recepting macrophages and monocytes can prolong the long half-life of the complex. Furthermore, given one the natural functions of Hp is to prevent cell-free Hb extravasation into tissue space (due to the large size of the Hp-Hb complex), apoHb-Hp-drug complexes should have very low rates of “leakage” into the tissue space.
As used herein, the term “tangential-flow filtration” refers to a process in which the fluid mixture containing the components to be separated by filtration is recirculated at high velocities tangential to the plane of the filtration membrane to reduce fouling of the filter. In such filtrations a pressure differential is applied along the length of the filtration membrane to cause the fluid and filterable solutes to flow through the membrane (i.e. filter). This filtration is suitably conducted as a batch process as well as a continuous-flow process. For example, the solution may be passed repeatedly over the membrane while that fluid which passes through the filter is continually drawn off into a separate unit or the solution is passed once over the membrane and the fluid passing through the filter is continually processed downstream.
As used herein, the term “ultrafiltration” is used for processes employing membranes rated for retaining solutes having a molecular weight between about 1 kDa and 1000 kDa.
As used herein, the term “reverse osmosis” refers to processes employing membranes capable of retaining solutes of a molecular weight less than 1 kDa such as salts and other low molecular weight solutes.
As used herein, the term “microfiltration” refers to processes employing membranes in the 0.1 to 10 micron pore size range.
As used herein, the expression “transmembrane pressure” or “TMP” refers to the pressure differential gradient that is applied along the length of a filtration membrane to cause fluid and filterable solutes to flow through the filter.
The term “hydrophobic,” as used herein, refers to a ligand which, as a separate entity, exhibits a higher solubility in a non-aqueous solution (e.g., octanol) than in water.
The term “conjugated protein,” as used herein, refers to a protein complex that includes an apoprotein and one or more associated hydrophobic ligands. The one or more hydrophobic ligands may by covalently or non-covalently associated with the apoprotein. Examples of conjugated proteins include, for example, lipoproteins, glycoproteins, phosphoproteins, hemoproteins, flavoproteins, metalloproteins, phytochromes, cytochromes, opsins, and chromoproteins.
The phrase “mild denaturing,” as used herein refers to a process which reversibly disrupts the secondary, tertiary, and/or quaternary structure of the conjugated protein, thereby facilitating separation of the hydrophobic ligand from the apoprotein. Mild denaturing can be distinguished from harsher conditions, which cleave the peptide backbone, primarily produce insoluble protein upon denaturation/renaturation, and/or disrupt protein structure to a degree such that the protein loses its biological function upon refolding.
The terms “isolating,” “purifying,” and “separating,” as used interchangeably herein, refer to increasing the degree of purity of a polypeptide or protein of interest or a target protein from a composition or sample comprising the polypeptide and one or more impurities (e.g., additional proteins or polypeptides).
Apohemoglobin-Haptoglobin Complexes
Provided herein are apohemoglobin-haptoglobin (apoHb-Hp) complexes.
In some embodiments, the apoHb-Hp complex can comprise apohemoglobin (apoHb) and haptoglobin (Hp) at a weight ratio of at least 1:1 (e.g., at least 1:1.1, at least 1:1.2, at least 1:1.3, at least 1:1.4, at least 1:1.5, at least 1:1.6, at least 1:1.7, at least 1:1.8, at least 1:1.9, at least 1:2, at least 1:2.1, at least 1:2.2, at least 1:2.3, at least 1:2.4, at least 1:2.5, at least 1:2.6, at least 1:2.7, at least 1:2.8, at least 1:2.9 or at least 1:3). In some embodiments, the apoHb-Hp complex can comprise apoHb and Hp at a weight ratio of 1:3 or less (e.g., 1:2.9 or less, 1:2.8 or less, 1:2.7 or less, 1:2.6 or less, 1:2.5 or less, 1:2.4 or less, 1:2.3 or less, 1:2.2 or less, 1:2.1 or less, 1:2 or less, 1:1.9 or less, 1:1.8 or less, 1:1.7 or less, 1:1.6 or less, 1:1.5 or less, 1:1.4 or less, 1:1.3 or less, 1:1.2 or less, or 1:1.1 or less).
The apoHb-Hp complex comprises apoHb and Hp at a weight ratio ranging from any of the minimum values described above to any of the maximum values described above. For example, in some embodiments, the apoHb-Hp complex can comprise apoHb and Hp at a weight ratio of from 1:1 to 1:3 (e.g., from 1:1.5 to 1:2.5, or from 1:1.7 to 1:2.2, or from 1:2.5 to 1:3).
In some embodiments, the Hp can be prepared using the ultrafiltration methods described below. For example, Hp can be prepared from plasma or fraction thereof (e.g., plasma fraction IV, plasma fraction V, a fraction of precipitated plasma (from salting out, or equivalent) or a combination thereof).
In certain embodiments, the Hp can have an average molecular weight of at least 70 kDa (e.g., at least 80 kDa, at least 90 kDa, at least 100 kDa, at least 150 kDa, at least 200 kDa, at least 250 kDa, at least 300 kDa, at least 350 kDa, at least 400 kDa, at least 450 kDa, at least 500 kDa, at least 550 kDa, at least 600 kDa, at least 650 kDa, at least 700 kDa, at least 750 kDa, at least 800 kDa, at least 850 kDa, at least 900 kDa, or at least 950 kDa). In certain embodiments, the Hp can have an average molecular weight of 1,000 kDa or less (e.g., 950 kDa or less, 900 kDa or less, 850 kDa or less, 800 kDa or less, 750 kDa or less, 700 kDa or less, 650 kDa or less, 600 kDa or less, 550 kDa or less, 500 kDa or less, 450 kDa or less, 400 kDa or less, 350 kDa or less, 300 kDa or less, 250 kDa or less, 200 kDa or less, 150 kDa or less, or 100 kDa or less).
The Hp can have an average molecular weight ranging from any of the minimum values described above to any of the maximum values described above. For example, in some embodiments, the Hp can have an average molecular weight of from 70 kDa to 1,000 kDa (e.g., from 80 kDa to 1,000 kDa, from 90 kDa to 800 kDa, from 80 kDa to 1,000 kDa, or from 80 kDa to 800 kDa).
In some embodiments, the apoHb can be prepared using the ultrafiltration methods described below. The apoHb prepared by various methods possess the same chemical identity (primary structure) and primarily the same quaternary conformation compared to apoHb prepared by existing precipitation or liquid-liquid extraction methodologies. The apoHb produced by the ultrafiltration methods described herein can exist in aqueous solution primarily as an αβ dimer without the use of reducing agents (2-mercaptoethanol, dithiothreitol). In contrast, previous methodologies may produce non-native tetramers (β2β2) that require reducing agents to form αβ dimers. Furthermore, the apoHb produced in the current methodology is stable for over a week at room temperature and stable at 4° C., −80° C. and in lyophilized form. Previous methodologies produced apoHb that quickly precipitated (approximately 24 hours) when stored at room temperature. In certain embodiments, the apoHb can be characterized by a residual Soret peak having a maximum absorption ranging from 411-417 nm, such as 412 nm (after renaturation/neutralization, but before complexation with Hp). Previous methodologies produced apoHb which had a residual Soret peak at 402-407 nm.
The apoHb-Hp complex can be formed by combining apoHb and Hp at an appropriate weight ratio. By way of example, since apoHb and Hp bind at a 1:1 molar ratio (apoHb αβ dimer binds to an αβ Hp dimer) that equates to 1:1 to 1:3 mass ratio depending on the Hp preparation and/or phenotype. ApoHb-Hp complexes can be formed by mixing apoHb and Hp at a weight ratio of at least 1:1 (e.g., at least 1:1.5, at least 1:2, or at least 1:3). By mixing an excess of apoHb with Hp, saturation of Hp Hb-binding sites can be achieved. Following complexation, the apoHb-Hp complex can be purified using tangential flow filtration (e.g., diafiltration using a 70 kDa TFF module to remove excess apoHb).
The apoHb and Hp can be wild-type proteins, recombinant proteins, or mutants. In certain embodiments, the apoHb can comprise an apoHb mutant which exhibits enhanced stability. Such mutants are known in the art, and described for example in U.S. Pat. No. 7,803,912 to Olson et al. which is incorporated herein by reference. In some examples, the apoHb can include one or more of the following amino acid mutations (the amino acids are specified by their helical location, i.e., A13 represents the thirteenth position along the A helix): α GlyA13 to Ala or Ser; α GlyB3 to Ala, Asp, Glu, or Asn; α CysG11 to Ser, Thr, or Val; β GlyA13 to Ala or Ser; β ProD2 to Ala; β GlyD7 to Lys; β GlyE13 to Ala, Thr, or Asp; βCysG14 to Val, Thr, Ser, or 11e; β ProH3 to Glu, Ala, or Gln; β CysG14 to Thr; β HisG18 to Ile, Leu, or Ala; β ProH3 to Glu; β TyrH8 to Trp or Leu; β ValH11 to Met, Leu, or Phe; or any combination thereof. Other apohemoglobins include, for example, α(H58L/V62F); β(H63L/V67F); αH87G; βH92G; βN108K; αV96W; and combinations thereof.
The apoHb-Hp complex can further include one or more active agents coordinated to the apoHb-Hp complex. In some cases, the active agent can be non-covalently associated with the apoHb-Hp complex. For example, in some cases, the active agent can be a hydrophobic active agent that non-covalently associates with the heme-binding region of apoHb. In other cases, the active agent can be covalently attached to the apoHb, covalently attached to the Hp, or a combination thereof.
In other embodiments, the active agent can be chemically linked (e.g., covalently bound) to an apohemoglobin-binding molecule such as heme. Such a system may present advantages for manufacturing as various drugs are hydrophobic which hinders their use in aqueous chemistry with proteins. Thus, the hydrophobic drugs could be chemically linked to an apohemoglobn-binding molecule, such as heme, in non-aqueous solvents. The linked active agent can then be solubilized via the binding of the apohemoglobin-binding molecules to the heme-binding pocket of apohemoglobin. In such embodiments, the active agent can be covalently tethered using the linking groups described below.
In certain embodiments, active agent can be covalently tethered to the apoHb, Hp, or a combination thereof via a linking group. When present, the linking group can be any suitable group or moiety which is at minimum bivalent, and connects the active agent to the protein. The linking group can be composed of any assembly of atoms, including oligomeric and polymeric chains. In some cases, the total number of atoms in the linking group can be from 3 to 200 atoms (e.g., from 3 to 150 atoms, from 3 to 100 atoms, from 3 and 50 atoms, from 3 to 25 atoms, from 3 to 15 atoms, or from 3 to 10 atoms).
In some embodiments, the linking group can be, for example, an alkyl, alkoxy, alkylaryl, alkylheteroaryl, alkylcycloalkyl, alkylheterocycloalkyl, alkylthio, alkylsulfinyl, alkyl sulfonyl, alkylamino, dialkylamino, alkylcarbonyl, alkoxycarbonyl, alkylaminocarbonyl, dialkylaminocarbonyl, or polyamino group. In some embodiments, the linking group can comprise one of the groups above joined to one or both of the moieties to which it is attached by a functional group. Examples of suitable functional groups include, for example, secondary amides (—CONH—), tertiary amides (—CONR—), secondary carbamates (—OCONH—; —NHCOO—), tertiary carbamates (—OCONR—; —NRCOO—), ureas (—NHCONH—; —NRCONH—; —NHCONR—, or —NRCONR—), carbinols (—CHOH—, —CROH—), ethers (—O—), and esters (—COO—, —CH2O2C—, CHRO2C—), wherein R is an alkyl group, an aryl group, or a heterocyclic group. For example, in some embodiments, the linking group can comprise an alkyl group (e.g., a C1-C12 alkyl group, a C1-C8 alkyl group, or a C1-C6 alkyl group) bound to one or both of the moieties to which it is attached via an ester (—COO—, —CH2O2C—, CHRO2C—), a secondary amide (—CONH—), or a tertiary amide (—CONR—), wherein R is an alkyl group, an aryl group, or a heterocyclic group. In certain embodiments, the linking group can be chosen from one of the following:
where m is an integer from 1 to 12 and R1 is, independently for each occurrence, hydrogen, an alkyl group, an aryl group, or a heterocyclic group.
If desired, the linker can serve to modify the solubility of the apoHb, Hp, and/or the apoHb-Hp complex. In some embodiments, the linker can be hydrophilic. In some embodiments, the linker can be an alkyl group, an alkylaryl group, an oligo- or polyalkylene oxide chain (e.g., an oligo- or polyethylene glycol chain), or an oligo- or poly(amino acid) chain.
In certain embodiments, the linker can be cleavable (e.g., cleavable by hydrolysis under physiological conditions, enzymatically cleavable, or a combination thereof). Examples of cleavable linkers include a hydrolysable linker, a pH cleavage linker, an enzyme cleavable linker, or disulfide bonds that are cleaved through reduction by free thiols and other reducing agents; peptide bonds that are cleaved through the action of proteases and peptidase; nucleic acid bonds cleaved through the action of nucleases; esters that are cleaved through hydrolysis either by enzymes or through the action of water in vivo; hydrazones, acetals, ketals, oximes, imine, aminals and similar groups that are cleaved through hydrolysis in the body; photo-cleavable bonds that are cleaved by the exposure to a specific wavelength of light; mechano-sensitive groups that are cleaved through the application of ultrasound or a mechanical strain (e.g., a mechanical strain created by a magnetic field on a magneto-responsive gel).
The active agent can comprise any suitable therapeutic or diagnostic agent.
In some embodiments, the therapeutic agent can comprise a diagnostic agent (e.g., an imaging agent, such as an MRI contrast agent). Suitable diagnostic agents can include molecules that are detectable in the body of a subject by an imaging technique such as X-ray radiography, ultrasound, computed tomography (CT), single-photon emission computed tomography (SPECT), magnetic resonance imaging (MRI), positron emission tomography (PET), optical fluorescent imaging, optical visible light imaging, and nuclear medicine including Cerenkov light imaging. For example, the diagnostic agent can comprise a radionuclide, paramagnetic metal ion, or a fluorophore.
In some cases, the diagnostic agent can comprise a metal chelator. The terms “metal chelator” and “chelating agent” refer to a polydentate ligand that can form a coordination complex with a metal atom. It is generally preferred that the coordination complex is stable under physiological conditions. That is, the metal will remain complexed to the chelator in vivo.
In some cases, the metal chelator is a molecule that complexes to a radionuclide metal or paramagnetic metal ion to form a metal complex that is stable under physiological conditions. The metal chelator may be any of the metal chelators known in the art for complexing a medically useful paramagnetic metal ion, or radionuclide.
In some cases, such as in the case of complexes designed for radiopharmaceutical or radiotherapy applications, it can be convenient to prepare the complexes comprising a radionuclide, at or near the site where they are to be used (e.g., in a hospital pharmacy or clinic). Accordingly, in some embodiments, the complex can comprise a metal chelator uncomplexed with a metal ion. In such embodiments, the complex can be complexed with a suitable metal ion prior to administration. In other embodiments, the complex comprises a metal chelator complexed with a suitable metal ion (e.g., a paramagnetic metal ion or a radionuclide).
Suitable metal chelators include, for example, linear, macrocyclic, terpyridine, and N3S, N2S2, or N4 chelators (see also, U.S. Pat. Nos. 4,647,447, 4,957,939, 4,963,344, 5,367,080, 5,364,613, 5,021,556, 5,075,099, 5,886,142, the disclosures of which are incorporated by reference herein in their entirety), and other chelators known in the art including, but not limited to, HYNIC, DTPA, EDTA, DOTA, TETA, and bisamino bisthiol (BAT) chelators (see also U.S. Pat. No. 5,720,934). For example, macrocyclic chelators, and in particular N4 chelators are described in U.S. Pat. Nos. 4,885,363; 5,846,519; 5,474,756; 6,143,274; 6,093,382; 5,608,110; 5,665,329; 5,656,254; and 5,688,487, the disclosures of which are incorporated by reference herein in their entirety. Certain N3S chelators are described in PCT/CA94/00395, PCT/CA94/00479, PCT/CA95/00249 and in U.S. Pat. Nos. 5,662,885; 5,976,495; and 5,780,006, the disclosures of which are incorporated by reference herein in their entirety. The chelator may also include derivatives of the chelating ligand mercapto-acetyl-glycyl-glycyl-glycine (MAG3), which contains an N3S, and N2S2 systems such as MAMA (monoamidemonoaminedithiols), DADS (N2S diaminedithiols), CODADS and the like. These ligand systems and a variety of others are described in Liu and Edwards, Chem. Rev. 1999, 99, 2235-2268; Caravan et al., Chem. Rev. 1999, 99, 2293-2352; and references therein, the disclosures of which are incorporated by reference herein in their entirety.
The metal chelator may also include complexes known as boronic acid adducts of technetium and rhenium dioximes, such as those described in U.S. Pat. Nos. 5,183,653; 5,387,409; and 5,118,797, the disclosures of which are incorporated by reference herein, in their entirety.
Examples of suitable chelators include, but are not limited to, derivatives of diethylenetriamine pentaacetic acid (DTPA), 1,4,7,10-tetraazacyclotetradecane-1,4,7,10-tetraacetic acid (DOTA), 1-substituted 1,4,7,-tricarboxymethyl 1,4,7,10 tetraazacyclododecane triacetic acid (DO3A), derivatives of the 1-1-(1-carboxy-3-(p-nitrophenyl)propyl-1,4,7,10 tetraazacyclododecane triacetate (PA-DOTA) and MeO-DOTA, ethylenediaminetetraacetic acid (EDTA), and 1,4,8,11-tetraazacyclotetradecane-1,4,8,11-tetraacetic acid (TETA), derivatives of 3,3,9,9-tetramethyl-4,8-diazaundecane-2,10-dione dioxime (PnAO); and derivatives of 3,3,9,9-tetramethyl-5-oxa-4,8-diazaundecane-2,10-dione dioxime (oxa PnAO). Additional chelating ligands are ethylenebis-(2-hydroxy-phenylglycine) (EHPG), and derivatives thereof, including 5-C1-EHPG, 5-Br-EHPG, 5-Me-EHPG, 5-t-Bu-EHPG, and 5-sec-Bu-EHPG; benzodiethylenetriamine pentaacetic acid (benzo-DTPA) and derivatives thereof, including dibenzo-DTPA, phenyl-DTPA, diphenyl-DTPA, benzyl-DTPA, and dibenzyl-DTPA; bis-2 (hydroxybenzyl)-ethylene-diaminediacetic acid (HBED) and derivatives thereof; the class of macrocyclic compounds which contain at least 3 carbon atoms and at least two heteroatoms (0 and/or N), which macrocyclic compounds can consist of one ring, or two or three rings joined together at the hetero ring elements, e.g., benzo-DOTA, dibenzo-DOTA, and benzo-NOTA, where NOTA is 1,4,7-triazacyclononane N,N′,N″-triacetic acid, benzo-TETA, benzo-DOTMA, where DOTMA is 1,4,7,10-tetraazacyclotetradecane-1,4,7,10-tetra(methyl tetraacetic acid), and benzo-TETMA, where TETMA is 1,4,8,11-tetraazacyclotetradecane-1,4,8,11-(methyl tetraacetic acid); derivatives of 1,3-propylenediaminetetraacetic acid (PDTA) and triethylenetetraaminehexaacetic acid (TTHA); derivatives of 1,5,10-N,N′,N″-tris(2,3-dihydroxybenzoyl)-tricatecholate (LICAM) and 1,3,5-N,N′,N″-tris(2,3-dihydroxybenzoyl)aminomethylbenzene (MECAM). Examples of representative chelators and chelating groups are described in WO 98/18496, WO 86/06605, WO 91/03200, WO 95/28179, WO 96/23526, WO 97/36619, PCT/US98/01473, PCT/US98/20182, and U.S. Pat. Nos. 4,899,755, 5,474,756, 5,846,519 and 6,143,274, each of which is hereby incorporated by reference in its entirety.
In some embodiments, the metal chelator comprises desferrioxamine (also referred to as deferoxamine, desferrioxamine B, desferoxamine B, DFO-B, DFOA, DFB or desferal) or a derivative thereof. See, for example U.S. Pat. Nos. 8,309,583, 4,684,482, and 5,268,165, each of which is hereby incorporated by reference in its entirety for its teaching of desferrioxamine and desferrioxamine derivatives.
As is well known in the art, metal chelators can be specific for particular metal ions. Suitable metal chelators can be selected for incorporation into the self-assembling molecule based on the desired metal ion and intended use of the self-assembling molecule.
Paramagnetic ions form a magnetic moment upon the application of an external magnetic field thereto. Magnetization is not retained in the absence of an externally applied magnetic field because thermal motion causes the spin of unpaired electrons to become randomly oriented in the absence of an external magnetic field. By taking advantage of its property of shortening the magnetic relaxation time of water molecules, a paramagnetic substance is usable as an active component of MRI contrast agents. Suitable paramagnetic transition metal ions include Cr3+, Co2+, Mn2+, Ni2+, Fe2+, Fe3+, Zr4+, Cu2+, and Cu3+. In preferred embodiments, the paramagnetic ion is a lanthanide ion (e.g., La3+, Gd3+, Ce3+, Tb3+, Pr3+, Dy3+, Nd3+, Ho3+, Pm3+, Er3+, Sm3+, Tm3+, Eu3+, Yb3+, or Lu3+). In MRI, especially preferred metal ions are Gd3+, Mn2+, Fe3+, and Eu2+.
MRI contrast agents can also be made with paramagnetic nitroxides molecules in place of the chelating agent and paramagnetic metal ion.
Suitable radionuclides include 99mTc, 67Ga, 68Ga, 66Ga, 47Sc, 51Cr, 167Tm, 141Ce, 111In, 123I, 125I, 131I, 124I, 18F, 11C, 15N, 17O, 168Yb, 175Yb, 140La, 90Y, 88Y, 86Y, 153Sm, 166Ho, 165Dy, 166Dy, 62Cu, 64Cu, 67Cu, 97Ru, 103Ru, 186Re, 188Re, 203Pb, 211Bi, 212Bi, 213Bi, 214Bi, 225Ac, 211At, 105Rh, 109Pd, 117mSn, 149Pm, 161Tb, 177Lu, 198Au, 199Au, 89Zr, and oxides or nitrides thereof. The choice of isotope will be determined based on the desired therapeutic or diagnostic application. For example, for diagnostic purposes (e.g., to diagnose and monitor therapeutic progress in primary tumors and metastases), suitable radionuclides includes 64Cu, 67Ga, 68Ga, 66Ga, 99mTc, and 111In, 18F, 89Zr, 123I, 124I, 177Lu, 15N, 17O. For therapeutic purposes (e.g., to provide radiotherapy for primary tumors and metastasis related to cancers of the prostate, breast, lung, etc.), suitable radionuclides include 64Cu, 90Y, 105Rh, 111In, 131I, 117 mSn, 149Pm, 153Sm, 161Tb, 166Dy, 166Ho, 175Yb, 177Lu, 186/188Re, 199Au, 131I, and 125I 212Bi, 211At.
In the case of complexes designed to be imaged using PET, radionuclides with short half-lives such as carbon-11 (˜20 min), nitrogen-13 (˜10 min), oxygen-15 (˜2 min), fluorine-18 (˜110 min), or rubidum-82 (˜1.27 min) are often used. In certain embodiments when a non-metal radionuclide is employed, the therapeutic or diagnostic agent comprises a radiotracer covalently attached to the self-assembling molecule. By way of exemplification, suitable 18F-based radiotracers include 18F-fluordesoxyglucose (FDG), 18F-dopamine, 18F-L-DOPA, 18F-fluorcholine, 18F-fluormethylethylcholin, and 18P-fluordihydrotestosteron.
In the case of self-assembled molecules designed to be imaged using PET, radionuclides with long half-lives such as 124I, or 89Zr are also often used.
Fluorescent imaging has emerged with unique capabilities for molecular cancer imaging. Fluorophores emit energy throughout the visible spectrum; however, the best spectrum for in vivo imaging is in the near-infrared (NIR) region (650 nm-900 nm). Unlike the visible light spectrum (400-650 nm), in the NIR region, light scattering decreases and photo absorption by hemoglobin and water diminishes, leading to deeper tissue penetration of light. Furthermore, tissue auto-fluorescence is low in the NIR spectra, which allows for a high signal to noise ratio. There is a range of small molecule organic fluorophores with excitation and emission spectra in the NIR region. Some, such as indocyanine green (ICG) and cyanine derivatives Cy5.5 and Cy7, have been used in imaging for a relatively long time. Modern fluorophores are developed by various biotechnology companies and include: Alexa dyes; IRDye dyes; VivoTag dyes and HylitePlus dyes. In general, the molecular weights of these fluorophores are below 1 kDa.
In some embodiments, the diagnostic agent can comprise a radiocontrast agent. In these embodiments, the diagnostic agent can comprise an iodinated moiety. Examples of suitable radiocontrast agents include iohexol, iodixanol and ioversol.
In some embodiments, the active agent can comprise a therapeutic agent. Any suitable therapeutic agent can be incorporated in the complexes described herein. In some examples, the therapeutic agent can comprise an agent to treat or prevent a disease or disorder associated with the overexpression of CD163. For example, in some cases, the therapeutic agent can comprise an anti-cancer agent, an anti-inflammatory agent, an agent that treats or prevents infection, or a combination thereof. In some examples, the active agent can comprise an agent administered to treat hemolytic anemia and other conditions characterized by or associated with hemolysis (e.g., sickle cell anemia, malaria, red blood cell transfusions, thalassemia, autoimmune disorders, bone marrow failure, infections, surgery, severe burns, acute lung injury, the administration of chemotherapeutics, radiation therapy, etc.). In some examples, the active agent can comprise an active agent administered to treat a disease associated with macrophages and monocytes. Such diseases are known in the art and include, for example, heart disease, HIV infection, cancer, fibrotic diseases (e.g., cystic fibrosis), asthma, inflammatory bowel disease, rheumatoid arthritis, and diseases in which macrophages or monocytes function as hosts for intracellular pathogens (e.g., malaria, tuberculosis, leishmaniasis, chikungunya, adenovirus, Legionnaires' disease, and infections caused by bacteria in the genus Brucella such as B. abortus, B. canis, B. melitensis, and B. suis).
In some embodiments, the active agent can comprise an anti-cancer agent. Examples of anti-cancer agents include, but are not limited to, Abiraterone Acetate, Abitrexate (Methotrexate), Abraxane (Paclitaxel Albumin-stabilized Nanoparticle Formulation), ABVD, ABVE, ABVE-PC, AC, AC-T, Adcetris (Brentuximab Vedotin), ADE, Ado-Trastuzumab Emtansine, Adriamycin (Doxorubicin Hydrochloride), Adrucil (Fluorouracil), Afatinib Dimaleate, Afinitor (Everolimus), Aldara (Imiquimod), Aldesleukin, Alemtuzumab, Alimta (Pemetrexed Disodium), Aloxi (Palonosetron Hydrochloride), Ambochlorin (Chlorambucil), Aminolevulinic Acid, Anastrozole, Aprepitant, Aredia (Pamidronate Disodium), Arimidex (Anastrozole), Aromasin (Exemestane), Arranon (Nelarabine), Arsenic Trioxide, Arzerra (Ofatumumab), Asparaginase Erwinia chrysanthemi, Avastin (Bevacizumab), Axitinib, Azacitidine, BEACOPP, Bendamustine Hydrochloride, BEP, Bevacizumab, Bexarotene, Bexxar (Tositumomab and I 131 Iodine Tositumomab), Bicalutamide, Bleomycin, Bortezomib, Bosulif (Bosutinib), Bosutinib, Brentuximab Vedotin, Busulfan, Busulfex (Busulfan), Cabazitaxel, Cabozantinib-S-Malate, CAF, Campath (Alemtuzumab), Camptosar (Irinotecan Hydrochloride), Capecitabine, CAPDX, Carboplatin, Carboplatin-Taxol, Carfilzomib, Casodex (Bicalutamide), CeeNU (Lomustine), Cerubidine (Daunorubicin Hydrochloride), Cervarix (Recombinant HPV Bivalent Vaccine), Cetuximab, Chlorambucil, Chlorambucil-Prednisone, CHOP, Cisplatin, Clafen (Cyclophosphamide), Clofarabine, Clofarex (Clofarabine), Clolar (Clofarabine), CMF, Cometriq (Cabozantinib-S-Malate), COPP, COPP-ABV, Cosmegen (Dactinomycin), Crizotinib, CVP, Cyclophosphamide, Cyfos (Ifosfamide), Cytarabine, Cytarabine (Liposomal), Cytosar-U (Cytarabine), Cytoxan (Cyclophosphamide), Dabrafenib, Dacarbazine, Dacogen (Decitabine), Dactinomycin, Dasatinib, Daunorubicin Hydrochloride, Decitabine, Degarelix, Denileukin Diftitox, Denosumab, DepoCyt (Liposomal Cytarabine), DepoFoam (Liposomal Cytarabine), Dexrazoxane Hydrochloride, Docetaxel, Doxil (Doxorubicin Hydrochloride Liposome), Doxorubicin Hydrochloride, Doxorubicin Hydrochloride Liposome, Dox-SL (Doxorubicin Hydrochloride Liposome), DTIC-Dome (Dacarbazine), Efudex (Fluorouracil), Elitek (Rasburicase), Ellence (Epirubicin Hydrochloride), Eloxatin (Oxaliplatin), Eltrombopag Olamine, Emend (Aprepitant), Enzalutamide, Epirubicin Hydrochloride, EPOCH, Erbitux (Cetuximab), Eribulin Mesylate, Erivedge (Vismodegib), Erlotinib Hydrochloride, Erwinaze (Asparaginase Envinia chrysanthemi), Etopophos (Etoposide Phosphate), Etoposide, Etoposide Phosphate, Evacet (Doxorubicin Hydrochloride Liposome), Everolimus, Evista (Raloxifene Hydrochloride), Exemestane, Fareston (Toremifene), Faslodex (Fulvestrant), FEC, Femara (Letrozole), Filgrastim, Fludara (Fludarabine Phosphate), Fludarabine Phosphate, Fluoroplex (Fluorouracil), Fluorouracil, Folex (Methotrexate), Folex PFS (Methotrexate), Folfiri, Folfiri-Bevacizumab, Folfiri-Cetuximab, Folfirinox, Folfox, Folotyn (Pralatrexate), FU-LV, Fulvestrant, Gardasil (Recombinant HPV Quadrivalent Vaccine), Gazyva (Obinutuzumab), Gefitinib, Gemcitabine Hydrochloride, Gemcitabine-Cisplatin, Gemcitabine-Oxaliplatin, Gemtuzumab Ozogamicin, Gemzar (Gemcitabine Hydrochloride), Gilotrif (Afatinib Dimaleate), Gleevec (Imatinib Mesylate), Glucarpidase, Goserelin Acetate, Halaven (Eribulin Mesylate), Herceptin (Trastuzumab), HPV Bivalent Vaccine (Recombinant), HPV Quadrivalent Vaccine (Recombinant), Hycamtin (Topotecan Hydrochloride), Hyper-CVAD, Ibritumomab Tiuxetan, Ibrutinib, ICE, Iclusig (Ponatinib Hydrochloride), Ifex (Ifosfamide), Ifosfamide, Ifosfamidum (Ifosfamide), Imatinib Mesylate, Imbruvica (Ibrutinib), Imiquimod, Inlyta (Axitinib), Intron A (Recombinant Interferon Alfa-2b), Iodine 131 Tositumomab and Tositumomab, Ipilimumab, Iressa (Gefitinib), Irinotecan Hydrochloride, Istodax (Romidepsin), Ixabepilone, Ixempra (Ixabepilone), Jakafi (Ruxolitinib Phosphate), Jevtana (Cabazitaxel), Kadcyla (Ado-Trastuzumab Emtansine), Keoxifene (Raloxifene Hydrochloride), Kepivance (Palifermin), Kyprolis (Carfilzomib), Lapatinib Ditosylate, Lenalidomide, Letrozole, Leucovorin Calcium, Leukeran (Chlorambucil), Leuprolide Acetate, Levulan (Aminolevulinic Acid), Linfolizin (Chlorambucil), LipoDox (Doxorubicin Hydrochloride Liposome), Liposomal Cytarabine, Lomustine, Lupron (Leuprolide Acetate), Lupron Depot (Leuprolide Acetate), Lupron Depot-Ped (Leuprolide Acetate), Lupron Depot-3 Month (Leuprolide Acetate), Lupron Depot-4 Month (Leuprolide Acetate), Marqibo (Vincristine Sulfate Liposome), Matulane (Procarbazine Hydrochloride), Mechlorethamine Hydrochloride, Megace (Megestrol Acetate), Megestrol Acetate, Mekinist (Trametinib), Mercaptopurine, Mesna, Mesnex (Mesna), Methazolastone (Temozolomide), Methotrexate, Methotrexate LPF (Methotrexate), Mexate (Methotrexate), Mexate-AQ (Methotrexate), Mitomycin C, Mitozytrex (Mitomycin C), MOPP, Mozobil (Plerixafor), Mustargen (Mechlorethamine Hydrochloride), Mutamycin (Mitomycin C), Myleran (Busulfan), Mylosar (Azacitidine), Mylotarg (Gemtuzumab Ozogamicin), Nanoparticle Paclitaxel (Paclitaxel Albumin-stabilized Nanoparticle Formulation), Navelbine (Vinorelbine Tartrate), Nelarabine, Neosar (Cyclophosphamide), Neupogen (Filgrastim), Nexavar (Sorafenib Tosylate), Nilotinib, Nolvadex (Tamoxifen Citrate), Nplate (Romiplostim), Obinutuzumab, Ofatumumab, Omacetaxine Mepesuccinate, Oncaspar (Pegaspargase), Ontak (Denileukin Diftitox), OEPA, OPPA, Oxaliplatin, Paclitaxel, Paclitaxel Albumin-stabilized Nanoparticle Formulation, Palifermin, Palonosetron Hydrochloride, Pamidronate Disodium, Panitumumab, Paraplat (Carboplatin), Paraplatin (Carboplatin), Pazopanib Hydrochloride, Pegaspargase, Peginterferon Alfa-2b, PEG-Intron (Peginterferon Alfa-2b), Pemetrexed Disodium, Perj eta (Pertuzumab), Pertuzumab, Platinol (Cisplatin), Platinol-AQ (Cisplatin), Plerixafor, Pomalidomide, Pomalyst (Pomalidomide), Ponatinib Hydrochloride, Pralatrexate, Prednisone, Procarbazine Hydrochloride, Proleukin (Aldesleukin), Prolia (Denosumab), Promacta (Eltrombopag Olamine), Provenge (Sipuleucel-T), Purinethol (Mercaptopurine), Radium 223 Dichloride, Raloxifene Hydrochloride, Rasburicase, R-CHOP, R-CVP, Recombinant HPV Bivalent Vaccine, Recombinant HPV Quadrivalent Vaccine, Recombinant Interferon Alfa-2b, Regorafenib, Revlimid (Lenalidomide), Rheumatrex (Methotrexate), Rituxan (Rituximab), Rituximab, Romidepsin, Romiplostim, Rubidomycin (Daunorubicin Hydrochloride), Ruxolitinib Phosphate, Sclerosol Intrapleural Aerosol (Talc), Sipuleucel-T, Sorafenib Tosylate, Sprycel (Dasatinib), Stanford V, Sterile Talc Powder (Talc), Steritalc (Talc), Stivarga (Regorafenib), Sunitinib Malate, Sutent (Sunitinib Malate), Sylatron (Peginterferon Alfa-2b), Synovir (Thalidomide), Synribo (Omacetaxine Mepesuccinate), Tafinlar (Dabrafenib), Talc, Tamoxifen Citrate, Tarabine PFS (Cytarabine), Tarceva (Erlotinib Hydrochloride), Targretin (Bexarotene), Tasigna (Nilotinib), Taxol (Paclitaxel), Taxotere (Docetaxel), Temodar (Temozolomide), Temozolomide, Temsirolimus, Thalidomide, Thalomid (Thalidomide), Toposar (Etoposide), Topotecan Hydrochloride, Toremifene, Torisel (Temsirolimus), Tositumomab and I131 Iodine Tositumomab, Totect (Dexrazoxane Hydrochloride), Trametinib, Trastuzumab, Treanda (Bendamustine Hydrochloride), Trisenox (Arsenic Trioxide), Tykerb (Lapatinib Ditosylate), Vandetanib, VAMP, Vectibix (Panitumumab), VeIP, Velban (Vinblastine Sulfate), Velcade (Bortezomib), Velsar (Vinblastine Sulfate), Vemurafenib, VePesid (Etoposide), Viadur (Leuprolide Acetate), Vidaza (Azacitidine), Vinblastine Sulfate, Vincasar PFS (Vincristine Sulfate), Vincristine Sulfate, Vincristine Sulfate Liposome, Vinorelbine Tartrate, Vismodegib, Voraxaze (Glucarpidase), Vorinostat, Votrient (Pazopanib Hydrochloride), Wellcovorin (Leucovorin Calcium), Xalkori (Crizotinib), Xeloda (Capecitabine), XELOX, Xgeva (Denosumab), Xofigo (Radium 223 Dichloride), Xtandi (Enzalutamide), Yervoy (Ipilimumab), Zaltrap (Ziv-Aflibercept), Zelboraf (Vemurafenib), Zevalin (Ibritumomab Tiuxetan), Zinecard (Dexrazoxane Hydrochloride), Ziv-Aflibercept, Zoladex (Goserelin Acetate), Zoledronic Acid, Zolinza (Vorinostat), Zometa (Zoledronic Acid), and Zytiga (Abiraterone Acetate). These anti-cancer agents are non-limiting, as the skilled artisan would be able to readily identify other anti-cancer agents.
In some embodiments, the active agent can comprise an anti-proliferative agent, e.g., mycophenolate mofetil (MMF), azathioprine, sirolimus, tacrolimus, paclitaxel, biolimus A9, novolimus, myolimus, zotarolimus, everolimus, or tranilast. These anti-proliferative agents are non-limiting, as the skilled artisan would be able to readily identify other anti-proliferative agents.
In some embodiments, the active agent can comprise an anti-inflammatory agent, e.g., corticosteroid anti-inflammatory drugs (e.g., beclomethasone, beclometasone, budesonide, flunisolide, fluticasone propionate, triamcinolone, methylprednisolone, prednisolone, or prednisone); or non-steroidal anti-inflammatory drugs (NSAIDs) (e.g., acetylsalicylic acid, diflunisal, salsalate, choline magnesium trisalicylate, ibuprofen, dexibuprofen, naproxen, fenoprofen, ketoprofen, dexketoprofen, fluribiprofen, oxaprozin, loxoprofen, indomethacin, tolmetin, sulindac, etodolac, ketorolac, diclofenac, aceclofenac, nabumetone, piroxicam, meloxicam, tenoxicam, droxicam, lornoxicam, isoxicam, mefenamic acid, meclofenamic acid, flufenamic acid, tolfenamic acid, celecoxib, rofecoxib, valdecoxib, parecoxib, lumiracoxib, etoricoxib, firocoxib, nimesulide, licofelone, H-harpaide, or lysine clonixinate). These anti-inflammatory agents are non-limiting, as the skilled artisan would be able to readily identify other anti-inflammatory agents.
In some embodiments, the active agent can comprise a drug that prevents or reduces transplant rejection, e.g., an immunosuppressant. Exemplary immunosuppressants include calcineurin inhibitors (e.g., cyclosporine, Tacrolimus (FK506)); mammalian target of rapamycin (mTOR) inhibitors (e.g., rapamycin, also known as Sirolimus); antiproliferative agents (e.g., azathioprine, mycophenolate mofetil, mycophenolate sodium); antibodies (e.g., basiliximab, daclizumab, muromonab); corticosteroids (e.g., prednisone). These drugs that prevent or reduce transplant rejection are non-limiting, as the skilled artisan would be able to readily identify other drugs that prevent or reduce transplant rejection.
In some embodiments, the active agent can comprise a drug that treats or prevents infection, e.g., an antibiotic. Suitable antibiotics include, but are not limited to, beta-lactam antibiotics (e.g., penicillins, cephalosporins, carbapenems), polymyxins, rifamycins, lipiarmycins, quinolones, sulfonamides, macrolides lincosamides, tetracyclines, aminoglycosides, cyclic lipopeptides (e.g., daptomycin), glycylcyclines (e.g., tigecycline), oxazonidinones (e.g., linezolid), and lipiarmycines (e.g., fidazomicin). For example, antibiotics include erythromycin, clindamycin, gentamycin, tetracycline, meclocycline, (sodium) sulfacetamide, benzoyl peroxide, and azelaic acid. Suitable penicillins include amoxicillin, ampicillin, bacampicillin, carbenicillin, cloxacillin, dicloxacillin, flucloxacillin, mezlocillin, nafcillin, oxacillin, penicillin g, penicillin v, piperacillin, pivampicillin, pivmecillinam, and ticarcillin. Exemplary cephalosporins include cefacetrile, cefadroxil, cephalexin, cefaloglycin, cefalonium, cefaloridine, cefalotin, cefapirin, cefatrizine, cefazaflur, cefazedone, cefazolin, cefradine, cefroxadine, ceftezole, cefaclor, cefamandole, cefmetazole, cefonicid, cefotetan, cefoxitin, cefprozil, cefuroxime, cefuzonam, cfcapene, cefdaloxime, cefdinir, cefditoren, cefetamet, cefixime, cefmenoxime, cefodizime, cefotaxime, cefpimizole, cefpodoxime, cefteram, ceftibuten, ceftiofur, ceftiolene, ceftizoxime, ceftriaxone, ceftazidime, cefclidine, cefepime, ceflurprenam, cefoselis, cefozopran, cefpirome, cequinome, ceftobiprole, ceftaroline, cefaclomezine, cefaloram, cefaparole, cefcanel, cefedrlor, cefempidone, cefetrizole, cefivitril, cefmatilen, cefmepidium, cefovecin, cefoxazole, cefrotil, cefsumide, cefuracetime, and ceftioxide.
Monobactams include aztreonam. Suitable carbapenems include imipenem/cilastatin, doripenem, meropenem, and ertapenem. Exemplary macrolides include azithromycin, erythromycin, larithromycin, dirithromycin, roxithromycin, and telithromycin. Lincosamides include clindamycin and lincomycin. Exemplary streptogramins include pristinamycin and quinupristin/dalfopristin. Suitable aminoglycoside antibiotics include amikacin, gentamycin, kanamycin, neomycin, netilmicin, paromomycin, streptomycin, and tobramycin. Exemplary quinolones include flumequine, nalidixic acid, oxolinic acid, piromidic acid, pipemidic acid, rosoxacin, ciprofloxacin, enoxacin, lomefloxacin, nadifloxacin, norfloxacin, ofoxacin, pefloxacin, rufloxacin, balofloxacin, gatifloxacin, repafloxacin, levofloxacin, moxifloxacin, pazufloxacin, sparfloxacin, temafloxacin, tosufloxacin, besifloxacin, clinafoxacin, gemifloxacin, sitafloxacin, trovafloxacin, and prulifloxacin. Suitable sulfonamides include sulfamethizole, sulfamethoxazole, and trimethoprim-sulfamethoxazone. Exemplary tetracyclines include demeclocycline, doxycycline, minocycline, oxytetracycline, tetracycline, and tigecycline. Other antibiotics include chloramphenicol, metronidazole, tinidazole, nitrofurantoin, vancomycin, teicoplanin, telavancin, linezolid, cycloserine, rifampin, rifabutin, rifapentin, bacitracin, polymyxin B, viomycin, and capreomycin. The skilled artisan could readily identify other antibiotics useful in the devices and methods described herein.
In some embodiments, the active agent can comprise an anti-HIV agent. Examples of anti-HIV agents include anti-HIV antibodies, immunostimulants such as interferon, and the like, a reverse transcriptase inhibitor, a protease inhibitor, an inhibitor of bond between a bond receptor (CD4, CXCR4, CCR5, and the like) of a host cell recognized by virus and the virus, and the like.
Specific examples of HIV reverse transcriptase inhibitors include Retrovir® (zidovudine or AZT), Epivir® (lamivudine or 3TC), Zerit® (sanilvudine), Videx® (didanosine), Hivid® (zalcitabine), Ziagen® (abacavir sulfate), Viramune® (nevirapine), Stocrin® (efavirenz), Rescriptor® (delavirdine mesylate), Combivir® (zidovudine+lamivudine), Trizivir® (abacavir sulfate+lamivudine+zidovudine), Coactinon® (emivirine), Phosphonovir®, Coviracil®, alovudine (3′-fluoro-3′-deoxythymidine), Thiovir (thiophosphonoformic acid), Capravirin (5-[(3,5-dichlorophenyl)thio]-4-isopropyl-1-(4-pyridylmethyl)imidazole-2-methanol carbamic acid), Tenofovir (PMPA), Tenofovir disoproxil fumarate ((R)-[[2-(6-amino-9H-purin-9-yl)-1-methylethoxy]methyl]phosphonic acid bis(isopropoxycarbonyloxymethyl)ester fumarate), DPC-083 ((4S)-6-chloro-4-[(1E)-cyclopropylethenyl]-3,4-dihydro-4-trifluoromethyl-2 (1H)-quinazolinone), DPC-961 ((4S)-6-chloro-4-(cyclopropylethynyl)-3,4-dihydro-4-(trifluoromethyl)-2 (1H)-quinazolinone), DAPD ((−)β-D-2,6-diaminopurine dioxolane), Immunocal, MSK-055, MSA-254, MSH-143, NV-01, TMC-120, DPC-817, GS-7340, TMC-125, SPD-754, D-A4FC, capravirine, UC-781, emtricitabine, alovudine, Phosphazid, UC-781, BCH-10618, DPC-083, Etravirine, BCH-13520, MIV-210, abacavir sulfate/lamivudine, GS-7340, GW-5634, GW-695634, and the like.
Specific examples of HIV protease inhibitors include Crixivan® (indinavir sulfate ethanolate), saquinavir, Invirase® (saquinavir mesylate), Norvir® (ritonavir), Viracept® (nelfinavir mesylate), lopinavir, Prozei® (amprenavir), Kaletra® (ritonavir+lopinavir), mozenavir dimesylate ([4R-(4α,5α,6β)]-1-3-bis[(3-aminophenyl)methyl]hexahydro-5,6-dihydroxy-4,7-bis(phenylmethyl)-2H-1,3-diazepin-2-one dimethanesulfonate), tipranavir (3′-[(1R)-1-[(6R)-5,6-dihydro-4-hydroxy-2-oxo-6-phenylethyl-6-propyl-2H-pyran-3-yl]propyl]-5-(trifluoromethyl)-2-pyridinesulfonamide), lasinavir (N-[5(S)-(tert-butoxycarbonylamino)-4(S)-hydroxy-6-phenyl-2(R)-(2,3,4-trimethoxybenzyl)hexanoyl]-L-valine 2-methoxyethylenamide), KNI-272 ((R)—N-tert-butyl-3-[(2S,3S)-2-hydroxy-3-N—[(R)-2-N-(isoquinolin-5-yloxyacetyl)amino-3-methylthiopropanoyl]amino-4-phenylbutanoyl]-5,5-dimethyl-1,3-thiazolidine-4-carboxamide), GW-433908, TMC-126, DPC-681, buckminsterfullerene, MK-944A (MK944 (N-(2(R)-hydroxy-1(S)-indanyl)-2(R)-phenylmethyl-4(S)-hydroxy-5-[4-(2-benzo[b]furanylmethyl)-2(S)-(tert-butylcarbamoyl)piperazin-1-yl]pentanamide)+indinavir sulfate), JE-2147 ([2(S)-oxo-4-phenylmethyl-3(5)-[(2-methyl-3-oxy)phenylcarbonylamino]-1-oxabutyl]-4-[(2-methylphenyl)methylamino]carbonyl-4(R)-5,5-dimethyl-1,3-thiazole), BMS-232632 ((3S,8S,9S,12S)-3,12-bis(1,1-dimethylethyl)-8-hydroxy-4,11-dioxo-9-(phenylmethyl)-6-[[4-(2-pyridinyl)phenyl]methyl]-2,5,6,10,13-pentaazatetradecanedicarboxylic acid dimethyl ester), DMP-850 ((4R,5S,6S,7R)-1-(3-amino-1H-indazol-5-ylmethyl)-4,7-dibenzyl-3-butyl-5,6-dihydroxyperhydro-1,3-diazepin-2-one), DMP-851, RO-0334649, Nar-DG-35, R-944, VX-385, TMC-114, Tipranavir, Fosamprenavir sodium, Fosamprenavir calcium, Darunavir, GW-0385, R-944, RO-033-4649, AG-1859, and the like.
The HIV integrase inhibitor may be S-1360, L-870810, and the like. The DNA polymerase inhibitor or DNA synthesis inhibitor may be Foscavir®, ACH-126443 (L-2′,3′-didehydro-dideoxy-5-fluorocytidine), entecavir ((1S,3S,4S)-9-[4-hydroxy-3-(hydroxymethyl)-2-methylenecyclopentyl]guanine), calanolide A ([10R-(10a,11(3,12a)]-11,12-dihydro-12-hydroxy-6,6,10,11-tetramethyl-4-propyl-2H,6H,10H-benzo[1,2-b:3,4-b′:5,6-b″]tripyran-2-one), calanolide B, NSC-674447 (1,1′-azobisformamide), Iscador (viscum alubm extract), Rubutecan, and the like. The HIV antisense drug may be HGTV-43, GEM-92, and the like. The anti-HIV antibody or other antibody may be NM-01, PRO-367, KD-247, Cytolin®, TNX-355 (CD4 antibody), AGT-1, PRO-140 (CCR5 antibody), Anti-CTLA-4 Mab, and the like. The HIV vaccine or other vaccine may be ALVAC®, AIDSVAX®, Remune®, HIV gp41 vaccine, HIV gp120 vaccine, HIV gp140 vaccine, HIV gp160 vaccine, HIV p17 vaccine, HIV p24 vaccine, HIV p55 vaccine, AlphaVax Vector System, canarypox gp160 vaccine, AntiTat, MVA-F6 Nef vaccine, HIV rev vaccine, C4-V3 peptide, p2249f, VIR-201, HGP-30W, TBC-3B, PARTICLE-3B, and the like, Antiferon (interferon-α vaccine), and the like.
The interferon or interferon agonist may be Sumiferon®, MultiFeron®, interferon-τ, Reticulose, Human leukocyte interferon alpha, and the like. The CCR5 antagonist may be SCH-351125, and the like. The pharmaceutical agent acting on HIV p24 may be GPG-NH2 (glycyl-prolyl-glycinamide), and the like. The HIV fusion inhibitor may be FP-21399 (1,4-bis[3-[(2,4-dichlorophenyl)carbonylamino]-2-oxo-5,8-disodium sulfonyl]naphthyl-2,5-dimethoxyphenyl-1,4-dihydrazone), T-1249, Synthetic Polymeric Construction No 3, pentafuside, FP-21399, PRO-542, Enfuvirtide, and the like. The IL-2 agonist or antagonist may be interleukin-2, Imunace®, Proleukin®, Multikine®, Ontak®, and the like. The TNF-α antagonist may be Thalomid® (thalidomide), Remicade® (infliximab), curdlan sulfate, and the like. The α-glucosidase inhibitor may be Bucast®, and the like.
The purine nucleoside phosphorylase inhibitor may be peldesine (2-amino-4-oxo-3H,5H-7-[(3-pyridyl)methyl]pyrrolo[3,2-d]pyrimidine), and the like. The apoptosis agonist or inhibitor may be Arkin Z®, Panavir®, Coenzyme Q10 (2-deca(3-methyl-2-butenylene)-5,6-dimethoxy-3-methyl-p-benzoquinone), and the like. The cholinesterase inhibitor may be Cognex®, and the like, and the immunomodulator may be Imunox®, Prokine®, Met-enkephalin (6-de-L-arginine-7-de-L-arginine-8-de-L-valinamide-adrenorphin), WF-10 (10-fold dilute tetrachlorodecaoxide solution), Perthon, PRO-542, SCH-D, UK-427857, AMD-070, AK-602, and the like.
In addition, Neurotropin®, Lidakol®, Ancer 20®, Ampligen®, Anticort®, Inactivin®, and the like, PRO-2000, Rev M10 gene, HIV specific cytotoxic T cell (CTL immunotherapy, ACTG protocol 080 therapy, CD4-ζ gene therapy), SCA binding protein, RBC-CD4 complex, Motexafin gadolinium, GEM-92, CNI-1493, (±)—FTC, Ushercell, D2S, BufferGel®, VivaGel®, Glyminox vaginal gel, sodium lauryl sulfate, 2F5, 2F5/2G12, VRX-496, Ad5gag2, BG-777, IGIV-C, BILR-255, and the like may be used in the combination therapy.
Other suitable active agents include porphyrin-based active agents (e.g., porphyrin-based imaging agents, porphyrin-based agents for photodynamic therapy), erythropoietin, hydroxycarbamide (also known as hydroxyurea), corticosteroids, immunosuppressive agents, analgesic agents, agents that induce hemolysis (e.g., rituximab, cephalosporins, dapsone, levodopa, levofloxacin, methyldopa, nitrofurantoin, NSAIDs, penicillin and derivatives thereof, phenazopyridine, quinidine), dexamethasone, conjugates targeting the CD163 receptor (e.g., agents described in U.S. Pat. No. 9,724,426 to Graversen et al. which is incorporated by reference in its entirety), antibiotics; anti-tuberculosis antibiotics (such as isoniazide, ethambutol); anti-retroviral drugs, for example inhibitors of reverse transcription (such as zidovudin) and/or protease inhibitors (such as indinavir); drugs with effect on leishmaniasis (such as meglumine antimoniate); immunosuppressive drugs such as a glucocorticoid (e.g., cortisone and derivatives thereof (such as hydrocortisone); prednisone and derivatives thereof (such as prednisolone, methylprednisolone, methylprednisolone-acetate, methylprednisolone-succinate); dexamethasone and derivatives thereof; triamcinolone and derivatives thereof (such as triamcinolonehexacetonuid, triamcinolonacetonamid); paramethasone; betamethasone; fluhydrocortisone; fluocinolone); methotrexate; cyclophosphamide; 6-mercaptopurin; cyclosporine; tacrolimus; mycophenolate mofetil; sirulimus; everolimus; an siRNA molecule capable of inhibiting synthesis of proinflammatory cytokines (such as TNF); a non-steroidal anti-inflammatory drug (NSAIDs, such as aspirin, ibuprofen); a steroid (such as vitamin D); and a disease-modifying anti-rheumatic drug (DMARDs, such as penicillamin, sulfasalazin, cyclosporine).
In some embodiments, the active agent can comprise a toll-like receptor (TLR) agonist. A “TLR agonist” as used herein, refers to a substance that can combine with a TLR and activate it. By slightly altering the structure of such substances, TLR agonists can be designed to have different stabilities in the body, allowing a certain amount of control over where the substances go, and how long they last. Microbial ligands have been identified for several mammalian TLRs. For example, TLR4 recognizes lipopolysaccharide (LPS), TLR2 interacts with peptidoglycan, bacterial lipopeptides, and certain types of LPS, TLR3 recognizes double-stranded RNA, TLR5 recognizes bacterial flagellin, TLR9 recognizes bacterial DNA.
TLR agonists are well-known in the art and include, for example, but not limited to, lipopolysaccharide (LPS, binds TLR4), Fibrin (binds TLR4), lipoteichoic acid (LTA, binds TLR2), peptidoglycan (PG, binds TLR2), CpG (bacterial DNA, binds TLR9), 7-thia-8-oxoguanosine (TOG or isatoribine, binds TLR7), 7-deazaguanosine (binds TLR7), 7-allyl-8-oxoguanosine (loxoribine, binds TLR7), 7-dezaguanosine (7-deza-G, binds TLR7), imiquimod (R837, binds TLR7), or R848 (binds TLR7). In some embodiments, the TLR agonist can comprise a TLR7 agonist or a TLR9 agonist that is carried by the apoHb-Hp complex for receptor mediated uptake and immune activation within the endosome. In certain embodiments, the TLR agonist can comprise a TLR7 agonist (e.g., an imidazoquinoline such as imiquimod).
The expression of heme oxygenase-1 (H0-1) inhibits vascular inflammation and the induction of apoptosis. Accordingly, in some embodiments, the active agent can comprise an agent which modulates the activity of heme-oxygenase-1 (H0-1) activity. In some embodiments, the modulator of HO-1 is an antagonist, partial agonist, inverse agonist, neutral or competitive antagonist, allosteric antagonist, and/or orthosteric antagonist of HO-1. In some embodiments, the modulator of HO-1 is a HO-1 agonist, partial agonist, and/or positive allosteric modulator. In some embodiments, the agonist, partial agonist, and/or positive allosteric modulator of HO-1 is piperine, hemin, and/or brazilin. In some embodiments, the active agent comprises a protoporphyrin IX complex, such as zinc protoporphyrin IX or tin protoporphyrin IX, that is a HO-1 antagonist.
Pharmaceutical Compositions
The complexes provided herein can be administered in the form of pharmaceutical compositions. These complexes can be prepared as described herein or elsewhere, and can be administered by a variety of routes, depending upon whether local or systemic treatment is desired and upon the area to be treated. Administration may be topical (including transdermal, epidermal, ophthalmic and to mucous membranes including intranasal, vaginal and rectal delivery), pulmonary (e.g., by inhalation or insufflation of powders or aerosols, including by nebulizer; intratracheal or intranasal), oral, or parenteral. Parenteral administration includes intravenous, intraarterial, subcutaneous, intraperitoneal, intramuscular or injection or infusion; or intracranial, (e.g., intrathecal or intraventricular, administration). Parenteral administration can be in the form of a single bolus dose, or may be, for example, by a continuous perfusion pump. In some embodiments, the complexes provided herein are suitable for parenteral administration. In some embodiments, the complexes provided herein are suitable for intravenous administration.
Pharmaceutical compositions and formulations for topical administration may include, but are not limited to, transdermal patches, ointments, lotions, creams, gels, drops, suppositories, sprays, liquids and powders. Conventional pharmaceutical carriers, aqueous, powder or oily bases, thickeners and the like may be necessary or desirable. In some embodiments, the pharmaceutical compositions provided herein are suitable for parenteral administration. In some embodiments, the pharmaceutical compositions provided herein are suitable for intravenous administration. In some embodiments, the pharmaceutical compositions provided herein are suitable for oral administration. In some embodiments, the pharmaceutical compositions provided herein are suitable for topical administration.
Also provided are pharmaceutical compositions which contain, as the active ingredient, a complex provided herein in combination with one or more pharmaceutically acceptable carriers (e.g. excipients). In making the pharmaceutical compositions provided herein, the complex can be mixed with an excipient, diluted by an excipient or enclosed within such a carrier in the form of, for example, a capsule, sachet, paper, or other container. When the excipient serves as a diluent, it can be a solid, semi-solid, or liquid material, which acts as a vehicle, carrier or medium for the active ingredient. Thus, the compositions can be, for example, in the form of tablets, pills, powders, lozenges, sachets, cachets, elixirs, suspensions, emulsions, solutions, syrups, aerosols (as a solid or in a liquid medium), ointments, soft and hard gelatin capsules, suppositories, sterile injectable solutions, and sterile packaged powders.
Some examples of suitable excipients include, without limitation, lactose, dextrose, sucrose, sorbitol, mannitol, starches, gum acacia, calcium phosphate, alginates, tragacanth, gelatin, calcium silicate, microcrystalline cellulose, polyvinylpyrrolidone, cellulose, water, syrup, and methyl cellulose. The formulations can additionally include, without limitation, lubricating agents such as talc, magnesium stearate, and mineral oil; wetting agents; emulsifying and suspending agents; preserving agents such as methyl- and propylhydroxy-benzoates; sweetening agents; flavoring agents, or combinations thereof.
The complexes can be effective over a wide dosage range and is generally administered in an effective amount. It will be understood, however, that the amount of the compound actually administered will usually be determined by a physician, according to the relevant circumstances, including the condition to be treated, the chosen route of administration, the actual compound administered, the age, weight, and response of the individual subject, the severity of the subject's symptoms, and the like.
The compositions provided herein can be administered one from one or more times per day to one or more times per week; including once every other day. The skilled artisan will appreciate that certain factors can influence the dosage and timing required to effectively treat a subject, including, but not limited to, the severity of the disease or disorder, previous treatments, the general health and/or age of the subject, and other diseases present. Moreover, treatment of a subject with a therapeutically effective amount of a complex described herein can include a single treatment or a series of treatments.
Dosage, toxicity and therapeutic efficacy of the complexes provided herein can be determined by standard pharmaceutical procedures in cell cultures or experimental animals, e.g., for determining the LD50 (the dose lethal to 50% of the population) and the ED50 (the dose therapeutically effective in 50% of the population). The dose ratio between toxic and therapeutic effects is the therapeutic index and it can be expressed as the ratio LD50/ED50. Complexes exhibiting high therapeutic indices are preferred.
In some embodiments, the composition can further comprise one or more additional peptides or proteins. In certain embodiments, the one or more additional proteins can comprise proteins that detoxify iron, detoxify heme, detoxify Hb or a combination thereof. For example, in some examples, the composition can further comprise transferrin, hemopexin, haptoglobin or a combination thereof. In some embodiments, the composition can further comprise additional (uncomplexed) apohemoglobin, additional (uncomplexed) haptoglobin, or a combination thereof
Methods of Use
The apoHb-Hp complexes described herein can be administered to subjects in need thereof to treat a variety of diseases and disorders.
The apoHb-Hp complexes described herein can be administered to a subject in need thereof, for example, to treat hemolytic anemia and other conditions characterized by or associated with hemolysis. Examples of such conditions include, for example, sickle cell anemia, thalassemia, hemoglobin C disease, hemoglobin SC disease, sickle thalassemia, hereditary spherocytosis, hereditary elliptocytosis, hereditary ovalcytosis, glucose-6-phosphate deficiency and other red blood cell enzyme deficiencies, paroxysmal nocturnal hemoglobinuria (PNH), paroxysmal cold hemoglobinuria (PCH), thrombotic thrombocytopenic purpura/hemolytic uremic syndrome (TTP/HUS), idiopathic autoimmune hemolytic anemia, drug-induced immune hemolytic anemia, secondary immune hemolytic anemia, non-immune hemolytic anemia caused by chemical or physical agents (e.g., chemotherapeutic agents, anti-infective agents), malaria, falciparum malaria, bartonellosis, babesiosis, clostridial infection, severe Haemophilus influenzae type b infection, extensive burns, transfusion reaction, rhabdomyolysis (myoglobinemia), transfusion of stored blood, cardiopulomonary bypass, hemodialysis, red blood cell transfusions, bone marrow failure, hemolytic anemia induced by infection, hemolytic anemia induced by surgery, acute lung injury, radiation-induced hemolytic anemia, and combinations thereof.
In some examples, the complexes described herein can be administered to treat hemolysis associated with sickle cell anemia, malaria, a red blood cell transfusion, thalassemia, an autoimmune disorder, bone marrow failure, an infection, a surgical procedure, a burn, an acute lung injury, sepsis, organ perfusion, the administration of a pharmaceutical agent, the administration of radiation therapy, and a combination thereof. In some examples, the complexes described herein can be administered prophylactically to a subject to prevent damage associated with anticipated hemolysis (e.g., prior to surgery, radiation therapy, acute radiation injury, etc.). In some examples, the complexes described herein can be co-administered with a therapy that induce hemolysis (e.g., a chemotherapeutic agent, an anti-infective agent, a radiation therapy, or a combination thereof).
The complexes described herein can also be added to compositions comprising red blood cells, for example, to stabilize these compositions.
The apoHb-Hp-active agent complexes described herein can also be used to target the delivery of drugs to macrophages or monocytes (e.g., to down-regulate production of inflammatory cytokines, to kill intracellular organisms, or to kill malignant cells). In this way, the complexes can be used to selectively deliver active agents that significant impact certain diseases while minimizing adverse impacts of the active agent on other cells in the body.
In some embodiments, compositions comprising an apoHb-Hp-active agent complex can be administered to a subject in need thereof to treat a disease characterized by the overexpression of CD163. Such diseases are known in the art, and include but not limited to, for example cancer (e.g., breast cancer, Hodgkin Lymphoma), liver cirrhosis, type 2 diabetes, macrophage activation syndrome, Gaucher's disease, sepsis, HIV infection, and rheumatoid arthritis.
In some embodiments, compositions comprising an apoHb-Hp-active agent complex can be administered to a subject in need thereof to treat a disease which involves macrophages or monocytes. Such diseases are known in the art and include, for example, heart disease, HIV infection, cancer, fibrotic diseases (e.g., cystic fibrosis), asthma, inflammatory bowel disease, rheumatoid arthritis, and diseases in which macrophages or monocytes function as hosts for intracellular pathogens (e.g., malaria, tuberculosis, leishmaniasis, chikungunya, adenovirus, Legionnaires' disease, coronavirus (e.g., SARS-CoV-2, SARS, MERS, etc.), and infections caused by bacteria in the genus Brucella such as B. abortus, B. canis, B. melitensis, and B. suis).
The apoHb-Hp complexes described herein (with or without associated active agents) can be administered to a subject in need thereof, for example, to treat hemolytic anemia and other conditions characterized by or associated with hemolysis. Examples of such conditions include, for example, sickle cell anemia, thalassemia, hemoglobin C disease, hemoglobin SC disease, sickle thalassemia, hereditary spherocytosis, hereditary elliptocytosis, hereditary ovalcytosis, glucose-6-phosphate deficiency and other red blood cell enzyme deficiencies, paroxysmal nocturnal hemoglobinuria (PNH), paroxysmal cold hemoglobinuria (PCH), thrombotic thrombocytopenic purpura/hemolytic uremic syndrome (TTP/HUS), idiopathic autoimmune hemolytic anemia, drug-induced immune hemolytic anemia, secondary immune hemolytic anemia, non-immune hemolytic anemia caused by chemical or physical agents (e.g., chemotherapeutic agents, anti-infective agents), malaria, falciparum malaria, bartonellosis, babesiosis, clostridial infection, severe Haemophilus influenzae type b infection, extensive burns, transfusion reaction, rhabdomyolysis (myoglobinemia), transfusion of aged blood, cardiopulomonary bypass, hemodialysis, red blood cell transfusions, bone marrow failure, hemolytic anemia induced by infection, hemolytic anemia induced by surgery, acute lung injury, radiation-induced hemolytic anemia, and combinations thereof.
In some examples, the complexes described herein can be administered to treat hemolysis associated with sickle cell anemia, malaria, a red blood cell transfusion, thalassemia, an autoimmune disorder, bone marrow failure, an infection, a surgical procedure, a burn, an acute lung injury, the administration of a pharmaceutical agent, the administration of radiation therapy, and a combination thereof. In some examples, the complexes described herein can be administered prophylactically to a subject to prevent damage associated with anticipated hemolysis (e.g., prior to surgery, radiation therapy, acute radiation injury, etc.). In some examples, the complexes described herein can be co-administered with a therapy that induce hemolysis (e.g., a chemotherapeutic agent, an anti-infective agent, a radiation therapy, or a combination thereof).
The complexes described herein can also be added to compositions comprising red blood cells, for example, to stabilize these compositions.
The apoHb-Hp complex may also be used to triger CD163+ uptake of drug-conjugated Hp. In addition to administering the full apoHb-Hp-drug complex, only the Hp-drug conjugate or apoHb-drug conjugate could be administered. Then, apoHb or Hp could be administered to induce macrophage and monocyte uptake of the complex. In doing so, the Hp-drug conjugate would have a longer circulatory half-life to perform its desired function. For example, similar to bispecific monoclonal antibodies, Hp could be complexed with a targeting agent and macrophage/monocyte uptake could be trigged by the injection of apoHb. Such a therapeutic approach could be employed to treat cancer. For cancer treatment, Hp conjugated with a cancer cell targeting molecule could be administered to a patient. The Hp conjugated with the cancer cell targeting molecule would circulate until it binds to the surface of the cancer cell. Subsequent administration of apoHb would bind to the Hp attached to the cancer cell. The resulting apoHb-Hp complex attached to the cancer cell would recruit macrophages and monocytes for phagocytosis of the cancerous cell.
In some embodiments, methods can further include administering an agent to a patient to modulate CD163 expression (and by extension circulation and/or delivery of the complexes described herein). For example, methods can comprise administering a gluticosteroid to the patient to increase expression of CD163 or administering an agent (e.g., a gene silencing agent) to decrease expression of CD163.
In some embodiments, the apoHb-Hp complex and an active agent coordinated thereto is administered in combination with an immunotherapy agent, such as an immune checkpoint inhibitor. In some examples, the immune checkpoint inhibitor can comprise an anti-PD1 or anti-PDL1 antibody. In some examples, the immune checkpoint inhibitor can comprise an anti-CTLA4 monoclonal antibody.
In some embodiments, the disease can involve cellular iron accumulation and ferroptosis. In some embodiments, the apoHb-Hp complex and the active agent (e.g., HO-1 enzyme agonist) coordinated thereto are administered in combination with a ferropototic agent, such as Bay117085 or withaferin A.
Methods of Purifying Apohemoglobin
Methods of isolating apoHb protein can comprise (i) contacting Hb with an aqueous solution comprising a water-miscible solvent and a pH modifier, thereby forming a protein solution having a pH of less than 6.5 or greater than 8; and (ii) filtering the protein solution by ultrafiltration against a filtration membrane having a pore size that separates the apohemoglobin from heme, thereby forming a retentate fraction comprising the apoHb and a permeate fraction comprising heme. In some embodiments, methods for isolating an apoHb can further comprise (iii) neutralizing the retentate fraction to isolate the apoHb.
In some examples, the Hb can be present in the protein solution at a concentration from 0.1 mg/mL to 5 mg/mL, such as from 0.5 mg/mL to 3 mg/mL.
The protein solution can have an acidic or basic pH, selected so as to facilitate dissociation of the hydrophobic ligand and the apoprotein.
In some cases, the protein solution can have an acidic pH. In some of these embodiments, the protein solution can have a pH of 6 or less (e.g., 5.5 or less, 5 or less, 4.5 or less, 4 or less, 3.5 or less, 3 or less, or 2.5 or less). In some embodiments, the protein solution can have a pH of 2 or more (e.g., 2.5 or more, 3 or more, 3.5 or more, 4 or more, 4.5 or more, 5 or more, or 5.5 or more).
The protein solution can have a pH ranging from any of the minimum values described above to any of the maximum values described above. For example, in some embodiments, the protein solution can have a pH of from 2 to 6, such as from 3 to 6.
In other cases, the protein solution can have a basic pH. In some of these embodiments, the protein solution can have a pH of greater than 8 (e.g., 8.5 or more, 9 or more, 9.5 or more, 10 or more, or 10.5 or more). In some embodiments, the protein solution can have a pH of 11 or less (e.g., 10.5 or less, 10 or less, 9.5 or less, 9 or less, or 8.5 or less.
The protein solution can have a pH ranging from any of the minimum values described above to any of the maximum values described above. For example, in some embodiments, the protein solution can have a pH of from greater than 8 to 11, such as from greater than 8 to 10.
Generally, the filtration membrane can be rated for retaining solutes having a molecular weight ranging from the molecular weight of the heme to the molecular weight of the apoHb (e.g., such as a membrane rated for retaining solutes having a molecular weight of from about 1 kDa to 100 kDa, such as from about 1 kDa to about 10 kDa).
In connection with the methods described herein, ultrafiltration can comprise direct-flow filtration (DFF), cross-flow or tangential-flow filtration (TFF), or a combination thereof. In certain embodiments, the ultrafiltration can comprise tangential-flow filtration (TFF).
The membranes useful in the filtration steps described herein can be in the form of flat sheets, rolled-up sheets, cylinders, concentric cylinders, ducts of various cross-section and other configurations, assembled singly or in groups, and connected in series or in parallel within the filtration unit. The apparatus can be constructed so that the filtering and filtrate chambers run the length of the membrane.
Suitable membranes include those that separate the desired species from undesirable species in the mixture without substantial clogging problems and at a rate sufficient for continuous operation of the system. Examples are described, for example, in Gabler FR. Tangential flow filtration for processing cells, proteins, and other biological components. ASM News 1984; 50:299-304. They can be synthetic membranes of either the microporous type or the ultrafiltration type. A microporous membrane has pore sizes typically from 0.1 to 10 micrometers, and can be made so that it retains all particles larger than the rated size. Ultrafiltration membranes have smaller pores and are characterized by the size of the protein that will be retained. They are available in increments from 1000 to 1,000,000 Dalton nominal molecular weight limits.
Generally, the filtration membrane can comprise an ultrafiltration membrane. Ultrafiltration membranes are normally asymmetrical with a thin film or skin on the upstream surface that is responsible for their separating power. They are commonly made of regenerated cellulose, polysulfone or polyethersulfone.
In some cases, each filtration step can involve filtration through a single filtration membrane. In other cases, because membrane filters are not perfect and may have pores that allow some intended retentate molecules to slip through, more than one membrane (e.g., two membranes, three membranes, four membranes, or more) having the same pore size can be utilized for a given filtration step. In these embodiments, the membranes can be placed so as to be layered parallel to each other (e.g., one on top of the other) such that filtered fluid sequentially flows through each of the more than one membrane.
Membrane filters for tangential-flow filtration are available as units of different configurations depending on the volumes of liquid to be handled, and in a variety of pore sizes. Particularly suitable for use in the methods described herein, on a relatively large scale, are those known, commercially available tangential-flow filtration units.
The filtration unit useful herein is suitably any unit now known or discovered in the future that serves as an appropriate filtration module, particularly for microfiltration and ultrafiltration. The preferred filtration unit is hollow fibers or a flat sheet device. These sandwiched filtration units can be stacked to form a composite cell. One example type of rectangular filtration plate type cell is available from Filtron Technology Corporation, Northborough, Mass., under the trade name Centrasette. Another example filtration unit is the Millipore Pellicon ultrafiltration system available from Millipore, Bedford, Mass.
The water-miscible solvent can comprise a polar protic solvent. In some embodiments, the water-miscible solvent can comprise an alcohol (e.g., ethanol, methanol, or a combination thereof).
In some embodiments, the aqueous solution can comprise at least 10% by volume (e.g., at least 15% by volume, at least 20% by volume, at least 25% by volume, at least 30% by volume, at least 35% by volume, at least 40% by volume, at least 45% by volume, at least 50% by volume, at least 55% by volume, at least 60% by volume, at least 65% by volume, at least 70% by volume, at least 75% by volume, at least 80% by volume, or at least 85% by volume) alcohol. In some embodiments, the aqueous solution can comprise 90% by volume or less (e.g., 85% by volume or less, 80% by volume or less, 75% by volume or less, 70% by volume or less, 65% by volume or less, 60% by volume or less, 55% by volume or less, 50% by volume or less, 45% by volume or less, 40% by volume or less, 35% by volume or less, 30% by volume or less, 25% by volume or less, 20% by volume or less, or 15% by volume or less) alcohol.
The aqueous solution can comprise a quantity of alcohol ranging from any of the minimum values described above to any of the maximum values described above. For example, in some embodiments, the aqueous solution can comprise from 60% to 90% by volume alcohol (e.g., 60% to 90% by volume ethanol).
In some embodiments, filtering step (ii) can comprise buffer exchange. In certain embodiments filtering step (ii) can comprise continuous diafiltration or dialysis.
Optionally, the retentate fraction can spectroscopically monitored during the continuous diafiltration to monitor separation of the heme from the apoHb. Spectroscopically monitoring the retentate fraction can comprise monitoring a spectroscopic peak (e.g., an absorbance peak) associated with the apoHb and a spectroscopic peak (e.g., an absorbance peak) associated with the Hb. In some embodiments, filtering step (ii) can comprise performing the continuous diafiltration until a relative magnitude of the absorbance peak associated with the apoHb and the absorbance peak associated with the Hb suggest that the apoHb and the Hb are present in the retentate fraction at a molar ratio of at least 9:1 (e.g., at least 10:1, at least 15:1, at least 20:1, at least 25:1, at least 50:1, or at least 100:1).
Methods for isolating an apoHb can further comprise (iii) neutralizing the retentate fraction to isolate the apoHb. In some embodiments, neutralizing step (iii) comprises continuous diafiltration with a buffer solution having a pH of from 6.8 to 7.6.
The purity of isolated apoHb can be assessed using a variety of methods known in the art, including for example, liquid chromatography and/or spectroscopic methods (UV-Vis spectroscopy, fluorescence spectroscopy, etc.). In certain embodiments, the apoHb isolated in step (iii) can comprise less than 1% (e.g., less than 0.75%, less than 0.5%, less than 0.25%, or less than 0.1%) residual heme relative to the concentration of apoHb isolated in step (iii), as measured by a suitable spectroscopic method (e.g., UV Vis spectroscopy).
The apoHb isolated in step (iii) can exhibit excellent stability relative to apoHb isolated using other conventional methodologies. In some embodiments, the apoHb isolated in step (iii) can be stable for a period of at least 7 days (e.g., at least 14 days, at least 30 days, at least 60 days, at least 120 days, or at least 180 days) at 22° C. In certain embodiments, at least 75% (e.g., at least 80%, at least 85%, at least 90%, or at least 95%) of the apoHb remains soluble in solution after storage at 22° C. for 7 days. In certain embodiments, at least 75% (e.g., at least 80%, at least 85%, at least 90%, or at least 95%) of the apoHb remains soluble in solution after storage at 4° C. for 180 days. In certain embodiments, at least 75% (e.g., at least 80%, at least 85%, at least 90%, or at least 95%) of the apoHb remains soluble in solution after storage at −80° C. for 180 days.
In some of these embodiments, at least 65% (e.g., at least 70%, at least 75%, at least 80%, or at least 85%) of the apoHb can retain its activity (i.e., retain its ability to bind heme) after storage at 22° C. for 7 days. In some of these embodiments, at least 65% (e.g., at least 70%, at least 75%, at least 80%, or at least 85%) of the apoHb can retain its activity (i.e., retain its ability to bind heme) after storage at 4° C. for 180 days. In some of these embodiments, at least 65% (e.g., at least 70%, at least 75%, at least 80%, or at least 85%) of the apoHb can retain its activity (i.e., retain its ability to bind heme) after storage at −80° C. for 180 days.
In some embodiments, methods can further comprise lyophilizing the apoHb isolated in step (iii).
Also described are methods of isolating apoHb from a protein solution comprising a Hb. These methods can comprise (i) mildly denaturing the Hb to form a protein solution; and (ii) filtering the protein solution by ultrafiltration against a filtration membrane having a pore size that separates the apoHb from the heme, thereby forming a retentate fraction comprising the apoHb and a permeate fraction comprising the heme. In some embodiments, methods for isolating an apoHb can further comprise (iii) neutralizing the retentate fraction to isolate the apoHb.
In some examples, mildly denaturing the Hb can comprise heating the Hb (e.g., to a temperature of from 40° C. to 60° C.).
In some examples, mildly denaturing the Hb can comprise contacting the Hb with a pH modifier (e.g., with an acid and/or a base). Mildly denaturing the Hb can comprise contacting the Hb with an effective amount of a pH modifier to produce an acidic or basic pH, selected so as to facilitate dissociation of the heme and the apoHb.
In some cases, mildly denaturing the Hb can comprise contacting the Hb with an effective amount of a pH modifier to produce a pH of 6 or less (e.g., 5.5 or less, 5 or less, 4.5 or less, 4 or less, 3.5 or less, 3 or less, or 2.5 or less). In some embodiments, mildly denaturing the Hb can comprise contacting the Hb with an effective amount of a pH modifier to produce a pH of 2 or more (e.g., 2.5 or more, 3 or more, 3.5 or more, 4 or more, 4.5 or more, 5 or more, or 5.5 or more).
Mildly denaturing the Hb can comprise contacting the Hb with an effective amount of a pH modifier to produce a pH ranging from any of the minimum values described above to any of the maximum values described above. For example, in some embodiments, mildly denaturing the Hb can comprise contacting the Hb with an effective amount of a pH modifier to produce a pH of from 2 to 6, such as from 3 to 6.
In other cases, mildly denaturing the Hb can comprise contacting the Hb with an effective amount of a pH modifier to produce a pH of 8 or more (e.g., 8.5 or more, 9 or more, 9.5 or more, 10 or more, or 10.5 or more). In some embodiments, mildly denaturing the Hb can comprise contacting the Hb with an effective amount of a pH modifier to produce a pH of 11 or less (e.g., 10.5 or less, 10 or less, 9.5 or less, 9 or less, or 8.5 or less.
Mildly denaturing the Hb can comprise contacting the Hb with an effective amount of a pH modifier to produce a pH ranging from any of the minimum values described above to any of the maximum values described above. For example, in some embodiments, mildly denaturing the Hb can comprise contacting the Hb with an effective amount of a pH modifier to produce a pH of from 8 to 11, such as from 8 to 10.
In some examples, mildly denaturing the conjugated protein can comprise contacting the Hb with a non-aqueous solvent, such as an alcohol. Examples of such non-aqueous solvents include, for example, ethanol, methanol, isopropanol, butanol, 2-propanol, phenol, or combinations thereof.
In some examples, mildly denaturing the Hb can comprise contacting the Hb with a chaotropic agent (e.g., a salt that can disrupt the structure of a protein by shielding charges and preventing the stabilization of salt bridges). Any salt in principle may be used. Examples of suitable chaotropic agents include, but are not limited to, guanidinium chloride, lithium perchlorate, lithium acetate, magnesium chloride, sodium dodecyl sulfate, thiourea, urea, calcium chloride, and combinations thereof.
ApoHb prepared by the filtration methods described herein (after renaturation/neutralization) can exhibit improved stability and purity as compared to apoHb prepared by existing precipitation and liquid-liquid extraction methodologies.
In some embodiments, the apoHb produced by the filtration methods described herein can be stable for a period of at least 7 days (e.g., at least 14 days, at least 30 days, at least 60 days, at least 120 days, or at least 180 days) at 22° C. In certain embodiments, at least 75% (e.g., at least 80%, at least 85%, at least 90%, or at least 95%) of the apoHb produced by the filtration methods described herein remains soluble in solution after incubation at 22° C. for 7 days. In certain embodiments, at least 75% (e.g., at least 80%, at least 85%, at least 90%, or at least 95%) of the apoHb produced by the filtration methods described herein remains soluble in solution after incubation at 4° C. for 180 days. In certain embodiments, at least 75% (e.g., at least 80%, at least 85%, at least 90%, or at least 95%) of the apoHb produced by the filtration methods described herein remains soluble in solution after incubation at −80° C. for 180 days.
In some of these embodiments, at least 65% of the apoHb can retain its activity (i.e., retain its ability to bind heme) after storage at 22° C. for 7 days. In some of these embodiments, at least 65% of the apoHb can retain its activity (i.e., retain its ability to bind heme) after storage at 4° C. for 180 days. In some of these embodiments, at least 65% of the apoHb can retain its activity (i.e., retain its ability to bind heme) after storage at −80° C. for 180 days.
The apoHb prepared by various methods possess the same chemical identity (primary structure) and primarily the same quaternary conformation compared to apoHb prepared by existing precipitation or liquid-liquid extraction methodologies. The apoHb produced by the ultrafiltration methods described herein can exist in aqueous solution primarily as an αβ dimer without the use of reducing agents (2-mercaptoethanol, dithiothreitol). In contrast, previous methodologies may produce non-native tetramers (a2(32) that require reducing agents to form αβ dimers. Furthermore, the apoHb produced in the current methodology is stable for over a week at room temperature and stable at 4° C., −80° C. and in lyophilized form. Previous methodologies produced apoHb that quickly precipitated (approximately 24 hours) when stored at room temperature. In certain embodiments, the apoHb can be characterized by a residual Soret peak having a maximum absorption ranging from 411-417 nm, such as 412 nm (after renaturation/neutralization, but before complexation with Hp).
Methods for the Purification of Haptoglobin
Haptoglobin (Hp) can be isolated from plasma or a fraction thereof. In some embodiments, methods for isolating Hp from plasma or a fraction thereof can comprise (i) clarifying the plasma or fraction thereof and (ii) filtering the clarified plasma or a fraction thereof by ultrafiltration against a filtration membrane, thereby forming a retentate fraction comprising Hp having a molecular weight of greater than about 100 kDa and a permeate fraction comprising serum proteins and other impurities having a molecular weight of less than about 100 kDa.
The plasma or fraction thereof can comprise plasma fraction IV, plasma fraction V, a fraction of precipitated plasma (from salting out, or equivalent) or a combination thereof.
Clarifying the plasma or a fraction thereof can comprise removing suspended solids from the plasma or fraction thereof. Removing suspended solids from the plasma or fraction thereof can comprise filtering (via ultrafiltration, microfiltration, depth filtration or equivalent) the plasma or a fraction thereof, contacting the plasma or a fraction thereof with a salting out agent (e.g., ammonium sulfate), an adsorbing agent (e.g., ethacridine lactate), or a combination thereof. Further clarification may be implemented through addition of a lipid-binding agent such as fumed silica (such as fumed silica sold under the tradename Aerosil380®, or similar), clay, bentonite, terra alba, active carbon, or a combination thereof.
In some embodiments, the ultrafiltration can comprise tangential-flow filtration.
In some cases, the method can further comprise filtering the permeate fraction comprising serum proteins and other impurities by ultrafiltration against a second filtration membrane, thereby forming a second retentate fraction comprising a blend of proteins having a molecular weight below about 100 kDa and above a cutoff value and a second permeate fraction comprising serum proteins and other impurities having a molecular weight below the cutoff value, wherein the blend of proteins in the second permeate comprises low molecular weight Hp, transferrin, hemopexin, or a combination thereof. The cutoff value can be from about 20 kDa to about 70 kDa, such as from about 25 kDa to about 50 kDa
In these methods, the second retentate fraction can include a blend of proteins (e.g., low molecular weight Hp, transferrin, hemopexin, or a combination thereof) that can bind and detoxify cell-free Hb, free iron, and/or free heme.
In other embodiments, methods for isolating Hp from plasma or a fraction thereof can comprise (i) filtering the plasma or fraction thereof by ultrafiltration against a first filtration membrane, thereby forming a first retentate fraction comprising serum proteins having a molecular weight above a first cutoff value and a first permeate fraction comprising most of the Hp and serum proteins having a molecular weight below the first cutoff value; and (ii) filtering the first permeate fraction by ultrafiltration against a second filtration membrane, thereby forming a second retentate fraction comprising small amounts of Hp2-1, Hp2-2, and serum proteins having a molecular weight below the first cutoff value and above a second cutoff value; and a second permeate fraction comprising Hp2-1, Hp2-2, and serum proteins having a molecular weight below the second cutoff value. In some cases, the method can further comprise (iii) filtering the second permeate fraction by tangential-flow filtration against a third filtration membrane, thereby forming a third retentate fraction comprising Hp2-1 and Hp2-2 having a molecular weight below the second cutoff value and above a third cutoff value; and a third permeate fraction comprising low molecular weight Hp, serum proteins and other impurities having a molecular weight below the third cutoff value. In some cases, the method can further comprise (iv) filtering the third permeate fraction comprising low molecular weight Hp, serum proteins and other impurities by ultrafiltration against a fourth filtration membrane, thereby forming a fourth retentate fraction comprising a blend of proteins having a molecular weight below the third cutoff value and above a fourth cutoff value and a fourth permeate fraction comprising serum proteins and other impurities having a molecular weight below the fourth cutoff value, wherein the blend of proteins in the retentate comprises low molecular weight Hp, transferrin, hemopexin, or a combination thereof.
The first cutoff value can be from about 650 kDa to about 1000 kDa. The second cutoff value can be from about 300 kDa to about 700 kDa. The third cutoff value can be from about 70 kDa to about 200 kDa. The fourth cutoff value can be from about 20 kDa to about 70 kDa. In certain examples, the first cutoff value can be about 750 kDa, the second cutoff value can be about 500 kDa, and the third cutoff value can be about 100 kDa. The fourth cutoff value can be about 30 kDa or about 50 kDa.
The plasma or fraction thereof can comprise plasma fraction IV, plasma fraction V, a fraction of precipitated plasma (from salting out, or equivalent) or a combination thereof.
Clarifying the plasma or a fraction thereof can comprise removing suspended solids from the plasma or fraction thereof. Removing suspended solids from the plasma or fraction thereof can comprise filtering (via ultrafiltration, microfiltration, depth filtration or equivalent) the plasma or a fraction thereof, contacting the plasma or a fraction thereof with a salting out agent (e.g., ammonium sulfate), an adsorbing agent (e.g., ethacridine lactate), or a combination thereof. Further clarification may be implemented through addition of a lipid-binding agent such as fumed silica (such as fumed silica sold under the tradename Aerosil380®, or similar), clay, bentonite, terra alba, active carbon, or a combination thereof.
In some embodiments, the ultrafiltration can comprise tangential-flow filtration.
In these methods, the fourth retentate fraction can include a blend of proteins (e.g., low molecular weight Hp, transferrin, hemopexin, or a combination thereof) that can bind and detoxify free Hb, free iron, and/or free heme.
Hp can also be isolated from a solution (e.g., plasma or a fraction thereof) by exploiting molecular size changes induced by protein complex formation. Such methods can comprise (i) filtering the protein solution by ultrafiltration against a first filtration membrane, thereby forming a first retentate fraction comprising impurities having a molecular weight above a first cutoff value and a first permeate fraction comprising the Hp and impurities having a molecular weight below the first cutoff value; (ii) contacting the first permeate fraction with a binding molecule that selectively associates with the Hp to form a Hp complex having a molecular weight above the first cutoff value; and (iii) filtering the first permeate fraction by ultrafiltration against a second filtration membrane (at the same or above the cut-off of the first membrane), thereby forming a second retentate fraction comprising the Hp complex having a molecular weight above the first cutoff value and a second permeate fraction comprising the impurities having a molecular weight below the first cutoff value.
In some cases, the Hp complex can be isolated (e.g., if the Hp itself is useful, or if the Hp complex is more stable under storage than the Hp or binding molecule). For example, in one example, the binding molecule can comprise apoHb (e.g., prepared as described above). In these embodiments, the resultant Hp complex can be an apoHb-Hp complex described herein.
In other cases, the method can further involve dissociating the Hp complex to re-form the Hp, and isolating the Hp. For example, the method can further comprise (iv) contacting the second retentate fraction with a dissociating agent, thereby inducing dissociation of the Hp complex to the Hp and the binding molecule, and (v) filtering the second retentate fraction to separate the Hp from the binding molecule and the dissociating agent, thereby isolating the Hp.
In some of these embodiments, step (v) can comprise filtering the second retentate fraction by ultrafiltration against a third filtration membrane, thereby forming a third retentate solution comprising the Hp having a molecular weight above a second cutoff value and a second permeate fraction comprising the impurities having a molecular weight below the second cutoff value.
If desired, ultrafiltration may be done with staging to improve separation between retained and filtrated solutes.
By way of non-limiting illustration, examples of certain embodiments of the present disclosure are given below.
The general schematic for the procedure to remove hydrophobic ligands from proteins employing the invention presented here is shown in
Apohemoglobin (apoHb) is a dimeric globular protein with two vacant heme-binding pockets that can bind heme or other hydrophobic ligands. Purification of apoHb is based on partial hemoglobin (Hb) unfolding to facilitate heme extraction into an organic solvent. However, current production methods are time consuming, difficult to scale up, and use highly flammable and toxic solvents. In this study, a novel and scalable apoHb production method was developed using an acidified ethanol solution to extract the hydrophobic heme ligand into solution and tangential flow filtration to separate heme from the resultant apoprotein. Total protein and active protein yields were >95% and ˜75%, respectively, with <1% residual heme in apoHb preparations and >99% purity from SDS-PAGE analysis. Virtually no loss of apoHb activity was detected at 4° C., −80° C., and in lyophilized form during long term storage. Structurally, size exclusion chromatography (SEC) and circular dichroism (CD) spectroscopy indicated that apoHb was dimeric with a ˜25% reduction of helical content compared to Hb. Furthermore, mass spectroscopy and reverse-phase chromatography indicated that the mass of the α and β subunits were virtually identical to the theoretical mass of these subunits in Hb and had no detectable oxidative modifications upon heme removal from Hb. SEC confirmed that apoHb bound to haptoglobin at similar ratio to that of native Hb. Finally, reconstituted Hb (rHb) was processed via a hemichrome removal method to isolate functional rHb for biophysical characterization in which the O2 equilibrium curve, O2 dissociation and CO association kinetics of rHb were virtually identical to native Hb. Overall, this study describes a novel and improved method to produce apoHb, as well as presents a comprehensive biochemical analysis of apoHb and rHb.
Human hemoglobin (Hb) is the major protein component contained inside human red blood cells (RBCs), and is well known for its role in oxygen (O2) storage and transport. It is a tetrameric protein (64 kDa), which consists of two pairs of αβ dimers (32 kDa) held together by non-covalent bonds. In each of the four globin chains (2a and 2(3 globins), a single heme prosthetic group is tightly bound inside the hydrophobic heme-binding pocket. Upon removal of heme from Hb, the resulting protein loses some of its helical content compared to native Hb. The resulting apoprotein is referred to as apohemoglobin (apoHb). ApoHb can react with heme to form reconstituted Hb (rHb), which shows virtually no difference in biophysical properties compared to native Hb. The heme-binding ability and heme-induced structural changes of apoHb make it an interesting precursor for studies into in vivo Hb synthesis and recombinant Hb production.
ApoHb is an attractive delivery vehicle for hydrophobic drug molecules, which can bind within the vacant heme-binding pockets. Heme is highly hydrophobic and cytotoxic; however, when bound inside the heme-binding pocket of Hb, its toxicity is reduced, and aqueous solubility increases. In addition to heme, other hydrophobic molecules such as modified hemes or therapeutic drug molecules can bind to the hydrophobic heme-binding pocket of apoHb.
Another exciting property of apoHb is its' clearance through CD163+ macrophages or monocytes. Similar to Hb, apoHb binds to haptoglobin (Hp). Hp is a plasma protein mainly responsible for the clearance of cell-free Hb. The apoHb-Hp/Hb-Hp complex is then recognized and uptaken by CD163+ macrophages and monocytes. This specific mode of clearance allows for targeted drug delivery to macrophages or monocytes. Thus, the ability of apoHb to bind hydrophobic molecules and facilitate targeted delivery towards CD163+ macrophages or monocytes make it a promising hydrophobic drug delivery vehicle.
From these properties, not only can apoHb be used as a drug carrier for hydrophobic molecules (i.e. molecules insoluble in aqueous solution) and targeted drug delivery to macrophages and monocytes, but its high heme affinity could be used to scavenge heme in vivo. States of hemolysis release cell-free Hb, which can lose its heme moiety. Free heme can undergo various redox-reactions, causing oxidation of various tissues. ApoHb could scavenge free heme, and thus forming cell-free Hb that can be cleared through CD163+ macrophages or monocytes. Furthermore, the hydrophobic molecule binding properties of apoHb can be used to bind MRI contrast agent molecules such as Mn-porphyrins (similar structure to normal Hb heme but with switching the Fe metal atom to Mn). The same idea applies to binding of fluorescent molecules to the vacant hydrophobic heme-binding pocket.
Another application of apoHb is its potential use in photodynamic therapy (PDT). PDT has recently been used to effectively treat cancers and other illnesses through the production of reactive oxygen species (ROS). The ROS produced by PDT surpasses the ability of cancer cells to resist cell apoptosis, disrupts the tumor vasculature and promotes shifting the immune system against the tumor. Furthermore, this treatment could be used for cancers like triple-negative breast cancer, in which commonly targeted receptors are not expressed. However, most photosensitizers (PS) lack specificity for tumor cells, have poor solubility, and cause systemic photosensitivity, inducing phototoxic and photoallergic reactions. Many PS targeting mechanisms are expensive, complicated to develop, or leak PS. Fortunately, the high expression of CD163+ tumor associated macrophages (TAM) in cancers could be targeted through their uptake of apohemoglobin (apoHb) via CD163 mediated endocytosis. PS bound to apoHb could improve PDT treatment by not only improving its biocompatibility and effectiveness, but also by specifically targeting a form of TAM that contributes to tumor growth. Furthermore, the metals in PS provide a second pathway for ROS production and anti-tumor immune response. Thus, treatment with PS bound apoHb could enhance the immunological shift against the tumor by lowering macrophage density and stimulating TAM differentiation to an anti-cancer phenotype. This immune change can destroy secondary tumors and prevent cancer metastasis and regression. We have successfully bound aluminum-phthalocyanines (a highly potent PS molecule currently undergoing clinical trials) to apoHb, thus increasing its' solubility in aqueous solution.
The first successful method for producing active apoHb was developed by Faneli et al. in 1958. In Fanelli's acetone extraction method, Hb was added to acidified acetone at −20° C. extracting heme into solution while precipitating the apoprotein (globin). After separation of the solid protein from the liquid phase, the apoprotein was re-dissolved in deionized (DI) water followed by extensive dialysis, yielding apoHb. Another procedure for active apoHb production was later developed by Teale in 1959, and further improved upon by Yonetani. In this process, heme was removed from Hb through exposure to acidified methylethylketone (MEK), forming two immiscible liquid layers. The heme partitions into the organic layer, while the globin partitions into the aqueous layer. After liquid-liquid separation of the layers, the aqueous globin solution underwent extensive dialysis similar to the acetone extraction method to yield apoHb.
However, the aforementioned apoHb production processes have various drawbacks that complicate scale up. First, both processes use highly flammable solvents and require costly separation equipment. Additionally, in the case of MEK extraction (the solvent most likely to be used for scale up), about 40% of the water-rich phase becomes saturated with MEK. Thus, extensive treatment is required to lower the MEK concentration in the aqueous phase before the aqueous phase can be safely discarded. The high MEK concentration in the aqueous phase will also require repeated MEK extractions to significantly lower the heme content of the purified apoHb. In the case of acetone extraction, not only is there an additional safety risk associated with centrifuging a flammable solvent, but the process may also require sequential acetone exposure to sufficiently remove heme, especially due to the possibility of heme entrapment within the protein precipitate. Finally, when acetone and MEK are used as heme extraction solvents at large-scales, buffer-exchange via dialysis will require large volumes of buffer and is a slow process. In this current work, a scalable and simple process for manufacturing apoHb is described.
Tangential flow filtration (TFF) is a size exclusion filtration technique greatly used in industrial biotechnology for purification of biomolecules due to its linear scalability, economic benefits and long membrane lifetime. Additionally, TFF easily facilitates controlled buffer exchange via diafiltration, which is preferable to extensive and lengthy dialysis and can reduce the equipment footprint for production. In this example, TFF was used to produce and purify apoHb using an acidic 80% ethanol solution (v/v) as the heme extraction solvent. It is important to note that ethanol poses a much lower flammability risk compared to previously used heme extraction solvents given that its flashpoint is 20° C. compared to −18° C. and −3° C. for acetone and MEK, respectively.
In this TFF process, the Hb precursor in aqueous solution was added to a TFF system filled with an acidic ethanol solution. Then, the mixture of acidic ethanol and Hb underwent continuous diafiltration in the TFF system with acidic ethanol as the diafiltration solution until sufficient heme was extracted from the mixture (i.e., the absorbance ratio of the Soret peak at 412 nm divided by the 280 nm protein peak was lower than 0.1). Next, DI water was used as the diafiltration solution to neutralize the acidic ethanol-Hb solution and to remove ethanol and any free heme from solution. Finally, the desired buffer for apoHb storage and analysis was used as the diafiltration solution for the last diafiltration step (a more detailed procedure for apoHb production via TFF can be found in the Methods Section).
Analysis of apoHb produced via the TFF purification process (TFF-apoHb) was accomplished by analyzing protein yields and biochemical properties of the resultant apoHb. The stability of apoHb as a function of storage time was also examined at different concentrations and temperatures via quantification of active and total protein of stored apoHb samples. The reconstituted Hb from TFF-apoHb also had its biophysical properties analyzed and compared to native Hb via its absorbance spectrum, O2 equilibrium curve, O2 dissociation and carbon monoxide (CO) binding kinetics.
Materials and Methods
Materials. Na2HPO4 (sodium phosphate dibasic), NaH2PO4 (sodium phosphate monobasic), NaHCO3(sodium bicarbonate), and hemin chloride were all procured from Sigma Aldrich (St. Louis, Mo.). KCN (potassium cyanide), HCl (hydrochloric acid), acetone, HPLC grade acetonitrile, HPLC grade trifluoroacetic acid (TFA), nylon syringe filters (rated pore size 0.22 μm), and dialysis tubing (rated pore size: 6-8 kDa) were purchased from Fisher Scientific (Pittsburgh, Pa.), while Millex-GP PES syringe filters (rated pore size: 0.2 μm) were purchased from Merck Millipore (Billerica, Mass.). Expired units of human RBCs were generously donated by the Transfusion Service in the Wexner Medical Center at The Ohio State University (Columbus, Ohio).
Hb Preparation. Human Hb for use in this study was prepared via TFF as described by Palmer et al. (Palmer, A. F., Sun, G. & Harris, D. R. Tangential flow filtration of hemoglobin. Biotechnol. Prog. 25, 189-199 (2009)). The concentration of Hb was determined spectrophotometrically based on the Winterbourn equation.
TFF-apoHb Preparation. A KrosFlo Research II TFF system (Spectrum Laboratories, Rancho Dominguez, Ca) with a single 10 kDa polysulfone (PS) hollow fiber (HF) module was used to purify apoHb from Hb. To examine the scalability of the purification process, the process was first performed on TFF filter (P/N: X11S-300-10S) with 20 cm2 surface area (MicroKros, Spectrum Laboratories, Rancho Dominguez, Ca) then scaled up to TFF filter (P/N: M11S-360-01S) with 1,050 cm2 surface area (MiniKros, Spectrum Laboratories, Rancho Dominguez, Ca). For both size filters, the individual HFs were 0.5 cm in diameter and were 20 cm in length. Purified Hb was added to a 80% (v/v) EtOH/DI water mixture containing 3 mM HCl (acidic ethanol) to achieve a maximum protein concentration of 2 mg/mL. For experiments with microKros filters, 18 mg of Hb was used as the basis. However, experiments utilizing miniKros filters (larger surface area than microKros filters) consisted of three batches with 1 g Hb, three batches with 1.2 g Hb and 8 batches with 2.0 g Hb (ran with two parallel HF modules) as the basis, respectively. The Hb dispersed in the acidic ethanol solution was continuously subjected to diafiltration with 9 times its' initial volume with acidic ethanol to remove heme from solution. After heme removal, the heme-free globin was subjected to diafiltration with DI water with 5 times its volume.
Finally, the apoHb solution was subjected to diafiltration with 5 times its initial volume using a final buffer solution consisting of either phosphate buffered saline (PBS, 10 mM phosphate, 137 mM NaCl, and 2.7 mM KCl, pH 7.4) or 0.1 M phosphate buffer (PB, pH 7.0). During processing, flow rates of 25 mL/min and 1.1 L/min were used for the microKros and miniKros HF modules, respectively. The transmembrane pressure was maintained at 7±1 psi with a back-pressure valve to facilitate optimal permeate flux. For large-scale production, a final concentration step was performed in which the apoHb solution volume was reduced to 50±10 mL then further concentrated on 10 kDa PS microKros filters to a final total protein concentration of 50-115 mg/mL. The entire process was performed in a cold room maintained at 4±1° C. After each run, TFF modules were rinsed with DI water followed by sanitization with 0.5 M NaOH. The modules were stored in 0.1 M NaOH and were extensively washed with DI water prior to use. A schematic of the apoHb TFF production schematic is shown in
Acetone ApoHb Preparation. The method commonly used to produce apoHb via acetone heme extraction was followed according to the protocol outlined by Fanelli et al.
Total Protein Assays. The total protein concentration of the apoHb solution was measured using a Coomassie Plus Protein assay kit (Pierce Biotechnology, Rockford, Ill.).
ApoHb Activity Assay. The activity of the heme-binding pocket of apoHb was determined via a dicyanohemin (DCNh) incorporation assay. Briefly, analysis of the equilibrium absorbance at 420 nm from apoHb and DCNh mixtures was used to determine the saturation point of apoHb heme-binding pockets with heme. The extinction coefficients of DCNh and rHbCN were 85 mM−1 cm−1 and 114 mM−1 cm−1, respectively.
ApoHb Stability. Three TFF-apoHb batches were prepared. Each batch was divided into three groups to test apoHb stability over time. These groups consisted of unconcentrated, concentrated and lyophilized apoHb. The unconcentrated group was obtained after buffer exchange of apoHb into PBS buffer at a concentration of −2 mg/mL. The remainder of the batch was either sent to be lyophilized or to be concentrated to −40 mg/mL. Immediately after production, apoHb activity and total protein was quantified via the DCNh activity assay and 280 nm absorbance, respectively. Of the concentrated and unconcentrated apoHb groups from each batch, samples were stored at either 37, 22, 4 or −80° C. for subsequent analysis. The lyophilized powder was stored in a closed container at −80° C. Additionally, the stored apoHb groups were reconstituted into rHb and had their absorbance spectra and O2 dissociation curve measured. After measuring the initial time point (immediately after production), each storage condition was assayed at varying time intervals to measure apoHb activity over time. These time intervals were chosen to capture relevant changes in activity at each storage condition. At 37° C., apoHb was expected to quickly lose activity, so measurements were made every 12 hours. In contrast, apoHb stored at −80° C. was expected to maintain activity for longer time durations. Thus, after initial measurements on a weekly basis, the insignificant changes lead to longer intervals between measurements. Statistical analysis was performed on JMP Pro v 12.2.2 (SAS Institute, Cary, N.C.) and measured concentrations were compared to the initial time point values of each batch. A linear fit with the logarithmic value of the concentration was used to examine the effect of time with the ANOVA test. For time point differences, time was considered as discrete and the TUKEY HSD test was performed.
Mass Spectroscopy. Before analysis, Hb and apoHb samples were buffer exchanged into 100 mM ammonium acetate (Fisher Scientific; San Jose, Calif.) using Micro Bio-Spin™ 6 columns (Bio-Rad; Hercules, Calif.). Samples were tested on a Finnigan LTQ mass spectrometer (Thermo Fisher Scientific, Waltham, Mass.) and analyzed using Xcalibur 2.2 software (Thermo Fisher Scientific, Waltham, Mass.). Samples from the same stock apoHb and Hb were then denatured in 1% acetic acid acetate (Fisher Scientific; San Jose, Calif.) and retested. The mass spectrometer parameters were: spray voltage: 1.5 kV; flow rate: 5 μL/min; capillary temperature: 200° C.; 3 microscans; and 100 ms injection time. The data was deconvoluted using mMass 5.5.0, (Copyright 2018 by Martin Strohalm).
Residual Heme Analysis. The residual heme in apoHb preparations was quantified via size exclusion chromatography (SEC). ApoHb samples prepared via TFF were separated on an analytical BioSep-SEC-53000 (600×7.5 mm) column (Phenomenex, Torrance, Calif.) attached to a Waters 2535 quaternary gradient module, Waters 2998 photodiode array multi-wavelength detector, and controlled using Empower Pro software (Waters Corp., Milford, Mass.). The mobile phase consisted of 50 mM potassium phosphate, pH 7.4. Since pigment-free proteins such as apoHb absorb at 280 nm and heme bound proteins such as Hb have a sharp Soret peak at 400-450 nm, the absorption wavelength was set at λ=280 nm to detect protein (although heme and heme-bound proteins also absorb at 280 nm), and λ=405 and 413 nm to detect protein containing heme. The number of heme molecules retained in apoHb preparations produced via TFF were determined by comparing the Soret spectra of four apoHb samples with the Soret spectra of a Hb sample of known concentration. The number of heme molecules in the apoHb preparation was then compared to the total protein of the sample (on a molar basis) to obtain the percentage of residual heme in each apoHb preparation.
Quaternary Structure. To estimate the quaternary structure of TFF-apoHb, apoHb and protein standards (conalbumin, 76 kDa; hHb, 64 kDa; carbonic anhydrase, 29 kDa; ribonuclease A, 14 kDa; and aprotinin, 6.5 kDa) were analyzed on a SEC column. The known molecular weight (MW) of the standards and their elution volumes were used to determine the coefficients (A, B) of a base 10 exponential function (MW=10A*(elution volume)+B) via non-linear regression. The estimated function parameters were used to estimate the MW of TFF-apoHb based on its elution volume. Samples were separated on an analytical Acclaim SEC-1000 (4.6×300 mm) column (Thermo Fisher Scientific, Waltham, Mass.) attached to a Dionex UltiMate 3000 system (Thermo Fisher Scientific, Waltham, Mass.). The mobile phase consisted of 50 mM potassium phosphate, pH 7.4. The flow rate and UV-visible spectral detection was controlled on Chromeleon 7 software with detection set to A=280 nm to detect protein elution at a flow rate 0.35 mL/min.
Haptoglobin Binding. To analyze haptoglobin (Hp) binding to TFF-apoHb, increasing concentrations of apoHb were mixed with haptoglobin (Hp) and the resultant mixture separated on a SEC column for analysis. Large molecular weight Hp (mixture of Hp2-2 and Hp2-1) was mixed with apoHb with a molecular weight of −32 kDa (dimeric apoHb) and separated on an analytical Acclaim SEC-1000 (4.6×300 mm) column (Thermo Fisher Scientific, Waltham, Mass.) attached to a Dionex UltiMate 3000 system (Thermo Fisher Scientific, Waltham, Mass.). The mobile phase consisted of 50 mM potassium phosphate, pH 7.4. The flow rate and UV-visible spectral detection was controlled on Chromeleon 7 software with detection set to A=280 nm to detect protein elution at a flow rate 0.35 mL/min. The percent change of the area under the curve between pure apoHb and a mixture of excess apoHb and Hp was used to determine the percent of apoHb that was bound to Hp. This percentage was compared to the mass of pure apoHb loaded to determine the Hp binding capacity of apoHb. The same procedure was repeated with Hb for comparison.
Reverse Phase Chromatography. Reverse phase (RP) chromatography was performed with a BioBasic-18 column (Thermo Scientific, Waltham, Mass.) on a Thermo Scientific Dionex Ultimate UHPLC system. The flow rate of the mobile phase was set to 0.75 mL/min. The column was equilibrated with 35% acetonitrile and 65% TFA (0.5 wt %, pH 2.6) for 10 minutes. The gradient was then shifted to 43% acetonitrile over a 1 minute interval. The protein was eluted with an increasing linear gradient of 43 to 47% acetonitrile for 30 minutes. The column was then held at 47% acetonitrile for 20 minutes.
Circular Dichroism. The far UV circular dichroism (CD) spectra of TFF-apoHb was measured on a JASCO J-815 CD spectrometer. Various TFF-apoHb samples and Hb were diluted in DI water to approximately 10 μM. The ellipticity of the samples was measured from 190-260 nm using a 0.1 mm path-length quartz cuvette. The change in alpha helical content of the apoglobin was determined via the ratio of the alpha-helix peak at 222 nm between TFF-apoHb to hHb.
Hb Reconstitution and Preparation. To regenerate the O2-binding capacity of Hb (i.e. reconstituted Hb, rHb), samples of apoHb were reconstituted with hematin to yield met-rHb and then reduced to yield rHb. First, hematin was added in excess to apoHb to yield met-rHb. The reaction was left overnight at 4±0.5° C. to go to completion. Met-rHb was centrifuged and passed through a 0.22 μm filter before any experiments were conducted. Reduction of met-rHb to yield deoxy-rHb was achieved by adding sodium dithionite at 1.5 mg/mL to met-rHb. The solution was then subjected to diafiltration on a 10 kDa TFF module to remove excess dithionite and any excess heme in solution using a modified HEMOX buffer (135 mM NaCl, 30 mM TES {N-[Tris (hydroxymethyl) methyl]-2-aminoethanesulfonic acid}, 5 mM KCl, pH 7.40±0.02 at 37° C.). During diafiltration, the system was open to the atmosphere to facilitate the conversion of rHb to oxygenated rHb.
Hemichrome Removal and rHb purification. rHb presented an altered absorbance spectra compared to pure Hb corresponding to the presence of hemichromes and/or heme bound to denatured globins. Two methods were developed to remove these unwanted species from solution. For analysis of small dilute samples in a spectrophotometer, passing the solution through a 0.22 μm nylon syringe filter removed the hemichromes by binding them to the hydrophobic filter membrane. However, this method was limited to processing small volumes of material, since the filter membrane would become saturated with these globin-heme species. When larger volumes of rHb were needed for analysis (i.e. more than 1 mg), an oxy-rHb sample was placed under a CO atmosphere to convert rHb into CO-rHb, a highly stable form of Hb. Then, the CO-rHb solution with the unwanted heme-globin complexes was heated to 65±1° C., and left under a CO atmosphere during about 100 minutes to precipitate hemichrome/heme. The hemichrome/heme precipitate was removed and the resulting CO-rHb solution was converted into oxy-rHb by placing it under a pure O2 stream for 2 hours.
rHb Analysis. Various liganded forms of rHb were analyzed via UV-Vis spectroscopy and compared against native Hb. The oxy-rHb and oxyHb equilibrium binding curves were measured using a Hemox analyzer (TCS Scientific Corp., New Hope, Pa.) at 37° C. The spectra of rHb was measured after one day reaction with excess heme (stage 1), after reduction and diafiltration with the modified HEMOX buffer (stage 2), after placing the rHb mixture under a CO atmosphere (stage 3), after heating the CO-rHb mixture and removing precipitate (stage 4), and after re-oxygenating the rHb sample (stage 5). Spectral deconvolution software was developed in the Python programming language (Python Software Foundation Beaverton, Oreg.) using the non-linear least squares function curve_fit of the SciPy package to determine the fraction of various liganded forms of rHb that contribute to the final spectra of the rHb mixture (i.e. containing metHb, hemichrome, oxyHb, HbCO and heme).
Stopped Flow Kinetics. CO binding to deoxyrHb, and 02 release from oxyrHb were measured using an Applied Photophysics SF-17 microvolume stopped-flow spectrophotometer (Applied Photophysics Ltd., Surrey, United Kingdom). Rapid kinetic measurements were performed using protocols previously described by Rameez and Palmer (Rameez, S. et al. Encapsulation of hemoglobin inside liposomes surface conjugated with poly(ethylene glycol) attenuates their reactions with gaseous ligands and regulates nitric oxide dependent vasodilation. Biotechnol. Prog. 28, 636-645 (2012)). Unmodified human Hb was used as a control. PBS (0.1 M, pH 7.4) was used as the reaction buffer for all kinetic measurements.
Results and Discussion
History of ApoHb Production Methods. The first published report of isolating globin (i.e., mixture of apoHb in its active and inactive forms) from Hb was from 1892 by Bertin-Sans and de Moitessier. In their method, oxygenated blood was coagulated with ether and mixed with a boiling solution of 10% tartaric acid in ethanol and further processed to yield apoHb. In 1898, Schulz made the first analysis of a purified globin solution produced via a mixture of ether and alcohol. Using this method, heme was extracted into the organic ether-ethanol phase leaving the globin in the aqueous phase. Later studies continued to use this protocol with slight modifications to produce globin until 1926. At that time, Hill and Holden noted that globin solutions were not soluble at the isoelectric point of Hb and the few available analyses of rHb were unclear on its biochemical properties. These issues were attributed to extensive globin denaturation (i.e., protein unfolding) from the use of harsh organic solvents at elevated temperatures, which lowered the yield of active apoglobin. Thus, Hill and Holden developed a very rigorous low temperature procedure that avoided the use of alcohol by using kieselguhr in ether to absorb heme. Using this method, Hill and Holden theorized it would not require protein unfolding, since it appeared that kieselguhr would remove heme under non-denaturing conditions. Yet, the theory of producing more active apoglobin due to the reduced protein unfolding step was shown to be highly improbable. Furthermore, Hsien Wu later noted that the major advancement in Hill and Holden's method was the low acidity of the solution and the low temperature of the process, and not because protein unfolding was minimized or abolished. It was also shown that performing Schulz's procedure under Hill and Holden's experimental conditions provided the same results as Hill and Holden. Finally, later studies by Ansos and Mirsky demonstrated the reversibility of protein unfolding, substantiating the idea that protein unfolding was not necessarily harsh for use in heme extraction. This idea of reversible protein unfolding led to the development of the acidic acetone heme extraction method, commonly used to this day. The acid-acetone procedure produces active soluble apoglobin even after unfolding the protein (to the point of precipitation) in acidic acetone.
Upon establishment of the previously mentioned acid-acetone or acid-methyl ethyl ketone (MEK) heme extraction procedures, these methods became the standard for producing apoHb in the literature. However, no modifications or improvements on these methods were made since their conception in the 1950s. These procedures require the use of highly flammable solvents which, when combined with the requirement of centrifugation or liquid-liquid extraction equipment, possess large safety risks for large-scale production. Additionally, since these processes use highly toxic solvents (acetone or MEK), it reduces possible biomedical applications of apoHb due to the presence of residual solvent in the preparation.
More recently developed apoglobin production methods employ acidified alcohols with subsequent separation facilitated by heme agglomeration, precipitation or adsorption on activated charcoal. Yet, these alcohol-derived apoglobins were made for applications in the food industry or for heme production, and did not provide an analysis of the retention of native apoHb activity or extent of heme removal. In this current study, the use of acidic ethanol-water heme extraction combined with TFF allows for the scalable and safe production of apoHb. A key advantage of this process is the absence of strong denaturants such as acetone or other ketones in the process. There is no method in the literature that describes purification of active apoHb from Hb in which both the heme and globin remain in the same phase. Additionally, previous apoHb production methods require extensive dialysis, which can be replaced by the quicker buffer exchange process facilitated by TFF run in diafiltration mode.
TFF Production of ApoHb. Earlier apoHb studies showed that the majority of purified apoHb product consisted of denatured apoHb with a small fraction of active apoHb in solution. Therefore, for applications where it is important to know the activity of the apoHb (i.e., defined as having a functional heme-binding pocket), apoHb quantification should be performed via an activity assay, and not total protein assays such as UV-absorbance analysis. Previous research quantified apoHb yield via analysis of the soluble apoglobin's absorbance peak at 280 nm (or via total protein assays). However, it has previously been demonstrated that quantification of soluble apoglobin does not accurately indicate the activity of the apoHb preparation, since a mixture of active and inactive apoglobins coexist in solution. As expected for a pigment-free protein, the UV-vis absorbance spectrum of apoHb consists of a single peak at 280 nm, which is shown in
Upon heme addition to apoHb, the absorbance of the solution at 280 nm and in the Soret band increases due to the presence of the heme pigment and from the covalent bond of the proximal histidine (His-F8) in apoHb with the heme iron. The final absorbance spectra of reconstituted Hb (rHb) should also be the same as native Hb with the characteristic intense Soret peak and Q-bands (discussed in rHb Analysis section). Since apoHb lacks this intense Soret band, heme extraction from Hb was determined through analysis of the UV-visible spectra of the apoHb solution. Successful heme extraction was determined when less than 1% residual heme could be detected (i.e., when the absorbance ratio between the protein peak at 280 nm to the Soret peak at ˜412 nm is less than 0.1).
The increase in Soret peak absorbance compared to pure heme when the porphyrin is incorporated into apoHb is due to the formation of a covalent bond between His-F8 in apoHb with the iron atom in heme. This difference in Soret peak absorbance is shown in
As shown in
After each batch, the DCNh-incorporation assay was used to quantify the moles of active apoHb on a per heme basis and the 280 nm absorbance (ε=12.7 mM−1 cm−1) was used to quantify moles of total protein. To determine protein yield, the moles of active apoHb and total protein were compared to the initial moles of heme in the Hb precursor. The yield of the TFF apoHb production method was compared to the acetone method using two size TFF filters (miniKros (surface area of 1,000 cm2) and microKros (surface area of 20 cm2)) to demonstrate the scalability of the TFF process. The results from this analysis are shown in Table 1.
ap < 0.05 compared with microKros TFF
b,cp < 0.05 between pairs with same letter
Studies have previously reported total apoHb yields from acetone or MEK extraction to be ˜90%. However, these studies quantified total protein (which includes both active and inactive apoHb) through methods such as protein absorbance at 280 nm. Thus, when comparing total protein yields, apoHb production via TFF (total protein yield of about 95%) had similar values to the commonly used heme extraction methods. Additionally, the total protein yield from acetone extraction agreed with previous reports. As expected, there was more total protein in solution compared to active apoHb in solution indicating that some of the resultant protein in solution lost its activity. It was observed that some protein was adsorbed on the filter membrane, which explains the loss in total protein. Yet, total protein analysis also showed that most of the protein was retained during production and that there was no significant difference between the studied production methods. Thus, since virtually no protein is lost in the TFF process, applications in which only heme-free globin is desired can still benefit from the TFF purification methodology. From the active protein analysis, acetone showed 70.3% yield compared to 83.1% and 73.4% for the microKros and miniKros TFF filters, respectively. Additionally, the yield from the small-scale miniKros filter was significantly different than both other setups. These results demonstrate that TFF production had similar or improved active apoHb yields compared to the acetone extraction method.
TFF-apoHb production with microKros filters had a significantly higher active apoHb yield than both the acetone and miniKros TFF methods (p<0.05). When scaling a TFF system, factors such as shear rate, pressure drop, filter type affect TFF efficiency. Therefore, operational parameters were kept constant when possible between the miniKros and microKros TFF systems in this example. However, the inlet pressure for the miniKros system prevented it from reaching the required flow rate to obtain the same shear rate as the microKros system. Thus, shear rates of 5,900 s−1 and 4,300 s−1, were achieved for the miniKros and microKros systems, respectively. The difference in shear rate could explain the lower permeate flow rate of the miniKros system, since lower shear rates may facilitate protein build up on the membrane. Since the permeate flow rate was not scaled, the diafiltration period was longer on the miniKros system, increasing the time that the protein remained unfolded in the acidified organic solvent. This longer exposure time could explain the lower active protein yield of the miniKros system compared to the microKros system and is a key variable which must be controlled to improve the yield of active apoHb.
During the concentration phase of TFF processing, protein precipitation was observed. Over time, HF membrane fouling decreased permeate flowrate up to 70%, making further concentration non-viable. To explore the effect of this concentration step on active protein yield and activity of the apoHb preparation, apoHb preparations were tested for total protein and active protein before and after concentration. Protein lost during concentration was compared to the initial mass of Hb used for apoHb production. As seen in Table 1, the loss of total protein from the sample was greater than active protein. Since more total protein was lost, the fraction of active protein in solution increased. The higher loss of inactive protein can be explained by the greater instability of inactive apoHb in solution versus active apoHb, facilitating precipitation at higher concentrations of apoHb. Stabilizing agents or alternative buffers may alter or improve these effects and should be considered in future method optimization.
The limit of 2 mg/mL of initial Hb precursor in the acidic ethanol solution was chosen to ensure full heme extraction from the sample. The dissociation of heme from unfolded globin seems to follow the equilibrium between the globin-heme complex and free heme+free globin in solution. Thus, when too high of an initial Hb concentration is loaded into the TFF circuit, the high globin concentration may limit the heme from dissociating from the globin-heme complex. Thus, in trials with a high initial Hb concentration little to no heme would permeate out of the TFF cartridge (undetectable on the absorbance spectrum). This was despite observing a significant amount of heme in the acidic ethanol solution within the TFF flow circuit (analyzed via the ratio of the Soret peak at 412 nm to the 280 nm protein peak). Additionally, the requirement of 9 diacycles for full heme extraction was evidenced by this equilibrium. If no heme retention occurred, the number of diacycles should have been closer to that of a simple buffer exchange (5 to 6 diacycles). Furthermore, when Hb solutions with concentrations of >50 mg/mL were used as the initial basis for the process, the protein rapidly denatured into a red precipitate when it made contact with the 80% acidic ethanol solution. Thus, to minimize this effect, Hb solutions with concentrations of −25 mg/mL were used when adding the holoprotein to the acidic ethanol solution.
Biophysical Properties of ApoHb produced via TFF. The TFF-apoHb was analyzed via electrospray ionization mass spectroscopy (ESI-MS) to determine if processing caused any amino acid residue modifications or protein damage.
Under native conditions, the observed mass of holo-Hb αβ dimers (
Further analysis of the quaternary structure of apoHb was performed using SEC. The HPLC-SEC profile of TFF-apoHb and human Hb is shown in
SEC was also performed on Hb and apoHb samples to analyze heme content (
A promising and important characteristic of apoHb is its clearance from the blood stream via CD163+ macrophage and monocyte mediated endocytosis. This in vivo clearance pathway for apoHb is the same for cell-free Hb. Both holo- and apo-Hb first bind serum Hp to form the (Hb/apoHb)-Hp complex, then the (Hb/apoHb)-Hp complex is captured by CD163+ macrophages and monocytes. To analyze if TFF-apoHb was capable of binding Hp, a fixed Hp concentration was mixed with increasing concentrations of apoHb and allowed to react to completion. The components of these mixtures were then separated via SEC-HPLC. As shown in
To ensure that no oxidative modifications or disulfide-bonded intermediates were present in our apoHb preparations, three different batches of TFF-apoHb were analyzed via reverse-phase HPLC (RP-HPLC). The samples were first evaluated via SEC-HPLC and, as shown in
RP-HPLC analysis is shown in
The secondary structure of TFF-apoHb was determined via CD of the far UV region (190-260 nm). This analysis is shown in
To test the activity of TFF-apoHb, the protein was reconstituted into rHb and the biophysical properties of rHb were analyzed and compared to native Hb. First, TFF-apoHb was reacted with heme and the spectral absorbance of various liganded forms of rHb were compared to native Hb. It was evident that the spectra of rHb in various liganded states shown in
Starting from a mixture of excess heme and apoHb, which reacts to form met-rHb, hemichromes, and excess heme (stage 1), the sample is processed to obtain oxy-rHb. Yet, hemichromes and excess non-specifically bound heme could be present in the sample along with oxy-rHb (stage 2). Thus, the mixture from stage 2 was placed under a CO atmosphere to transform oxy-rHb into the more heat stable CO-rHb species (stage 3). HbCO is more resistant to thermal denaturation and precipitation at elevated temperatures (65° C.) compared to other liganded forms of Hb, whereas heme-globin complexes and hemichromes are highly unstable and precipitate even at low (4° C.) temperatures. Thus, when a mixture of CO-rHb and heme-globin complexes is heated, the CO-rHb remains in solution while the unstable species precipitate out of solution (stage 4). After heating and separation of hemichromes, the CO-rHb can be reverted into oxy-rHb under a pure 02 atmosphere and white light illumination (stage 5).
To analyze this unique hemichrome removal method and provide better information on the species present in solution, a spectral deconvolution program was developed and implemented to determine the fraction of rHb species present in solution during the various processing stages in the reconstitution process. As shown in
To further analyze the biophysical properties of rHb derived from TFF-apoHb heme extraction, rHb was fully reconstituted back to oxy-rHb and its O2 equilibrium binding curve measured using a HEMOX Analyzer. From the O2 equilibrium curve, the P50 (partial pressure of O2 required to saturate half of the heme binding sites with O2) and cooperativity coefficient (n) can be regressed. The O2 dissociation (koff,O
Previous studies have shown that rHb produced from acetone extraction has the same biophysical properties compared to native Hb. This was confirmed from the quantitative results listed in Table 2. The P50 of rHb (11.36±0.87 and 11.20±0.43 mmHg for concentrated and unconcentrated rHb, respectively) was similar to native Hb (11.69±0.88 mmHg), with no statistical difference (p<0.05). The cooperativity of rHb from TFF-apoHb (2.14±0.17 and 2.27±0.10 for concentrated and unconcentrated rHb, respectively) was statistically different (p<0.05) compared to native Hb (2.73±0.11). Yet, similar to native Hb, TFF derived rHb had cooperativity greater than 2, which is indicative of cooperative O2 binding. Additionally, previous studies have shown that Hb which has been oxidized then reduced possess lower cooperativity than native Hb. This effect could have been exacerbated by the formation of reactive free radicals upon use of dithionite for reduction. It has also been shown that, upon reconstitution, the heme can enter the heme pocket in an altered orientation, which also reduces rHb cooperativity. Thus, the lower cooperativity of rHb compared to native Hb can be due to incorrect heme insertion and the necessary reduction step to form oxy-rHb. Also shown in Table 2 are the rate constants for O2 dissociation from HbO2 and CO association to deoxyHb. There was no statistical significant difference between the CO association rate constants for native Hb (kon,CO=180±7 nM/s) and TFF rHb (kon,CO=175±4 nM/s). However, the difference in O2 dissociation between native Hb (koff,O
ap < 0.05 compared with hHb
To analyze the stability of TFF-apoHb during storage, the amount of active and total apoHb was assessed under different storage conditions (37, 22, 4, −80° C. or lyophilized) and at two protein concentrations (concentrated [33.80±0.36 mg/mL active apoHb with 41.40±2.77 mg/mL total protein] or unconcentrated [1.47±0.01 mg/mL active apoHb with 1.99±0.17 mg/mL total protein]). The results from this analysis are shown in
At physiological core body temperature (37° C.) (
While stored at 4° C. (
Under all concentration dependent conditions, the concentrated samples had statistically significant lower active apoHb retention compared to unconcentrated samples, indicating that higher storage concentrations led to higher active apoHb loss. This observation can be explained by the higher probability of protein aggregation at higher protein concentrations. On the other hand, total protein retention was not statistically significant between concentrated and unconcentrated samples, except for the 37° C. storage condition. These observations indicated that the concentration of stored samples only significantly influenced apoHb activity. Furthermore, lyophilized apoHb and storage at 37° C. appear to have coinciding total protein and active protein retention, indicating that the protein lost from heating or from the freeze-drying process was not selective for whether the protein was active or not. To further test the biophysical properties of the apoHb samples stored at 4° C., samples were fully reconstituted into rHb after one month of storage and the P50 and cooperativity coefficient did not show any significant difference compared to apoHb samples reconstituted to rHb right after TFF production.
The stability of TFF-apoHb under different storage conditions was also assessed by RP-HPLC, CD and SEC-HPLC. This analysis is shown in
To further investigate the effects of prolonged storage on TFF-apoHb, the CD spectra of stored samples was measured to analyze any loss of alpha helical content. All samples were diluted in DI water to approximately 10 μM to remove any interference from salt. The far UV CD spectra was measured for the diluted samples, and these results are shown in
Given that the literature on apoHb indicates that the apoprotein is unstable at room temperature and that TFF-apoHb was shown to be relatively stable at 22° C., a TFF-apoHb sample that had been stored at 4° C. for over a year was left at 22° C. in a sealed cuvette to monitor protein loss via precipitation. The results from this experiment is shown in
An analysis of the potential tetramer-dimer equilibrium of the apoprotein was performed by testing samples that contained detectable tetramers on the SEC-HPLC. These results are shown in
Overall, given that there have been various reports in the literature on oligomerization of apoHb prepared via either acid-acetone or MEK methodologies, the formation of these non-native species may be linked to the preparation of apoHb. Not only was there minimal amounts of tetrameric species in our TFF-apoHb preparations, but there were no higher orders species detected under any of the tested storage conditions. Previous SEC-HPLC of MEK-apoHb showed a large percentage of tetrameric species and other higher order aggregates, which required the use of a reducing agent (such as dithiothreitol) during preparation to form dimeric apoHb. Furthermore, these tetrameric species were found to be dissociated at low concentrations, indicating that these species were not formed via irreversible disulfide bonds. Finally, even dimeric apoHb may also dissociate into monomers at low concentrations.
Conclusions
The possible biomedical applications of apoHb are very promising. Yet, its wide-scale use and analysis is restricted by current production methods, which are not easily scalable. New techniques to produce active apoHb have not been presented for decades even though new bioprocessing techniques have been developed. Existing apoHb production protocols require extensive dialysis and the use of highly flammable solvents. The newly proposed acidic alcohol TFF apoHb production method provides an easy and scalable method for producing active apoHb with more than 95% total protein and 75% active protein yields. Through the use of a 80% (v/v) ethanol:water solution for heme extraction, the flammability risks and toxicity issues with residual solvent are drastically lowered compared to previous apoHb production methods. Yet, the most valuable benefit of this new method is the use of an easily and cost-effective scalable process such as TFF for protein purification.
TFF-apoHb has the same characteristics as apoHb produced via previously published methodologies in the literature. These characteristics include: heme-binding activity, Hp binding activity, exists primarily as a dimeric species in aqueous solution, rHb O2 equilibria and ligand binding kinetics. Yet, unlike apoHb produced previously in the literature, stability studies showed that TFF-apoHb can be stored at 4° C., −80° C. and in lyophilized form without appreciable changes in activity with higher stability at room temperature compared to previous apoHb storage studies. Additionally, ESI-MS analysis of TFF-apoHb demonstrated that it retained its structure without any chemical modifications. RP-HPLC demonstrated that no oxidative modifications were present in freshly prepared TFF-apoHb nor for samples lyophilized or stored at −80° C. Quaternary structure analysis via SEC-HPLC showed that TFF-apoHb αβ dimers did not form appreciable amounts of tetramers over prolonged storage at 4° C. Furthermore, new insight into apoHb oxidation and degradation was provided based on the oxidation of the β-chain of apoHb when stored at 4° C. or 22° C. for prolonged periods of time. Evidence for both tetramer-dimer and dimer-monomer equilibrium of apoHb was also presented. Finally, an improved hemichrome removal procedure was developed that could generate rHb absorbance spectra indistinguishable from native Hb. Also, rHb generated from TFF-apoHb demonstrated that the reconstituted protein maintains native Hb-like O2 dissociation and CO association kinetics.
Taken together, this example presents an improved method for producing apoHb with a comprehensive analysis of the relevant biophysical properties apoHb and rHb.
The example describes a scalable process to enable purification of haptoglobin (Hp) from serum proteins in plasma or plasma fractions through the use of tangential flow filtration (TFF). The TFF process brackets Hp in a molecular weight range with low levels of common serum proteins, yielding a product that can include polymeric forms of Hp.
Haptoglobin (Hp) is an α-2 glycoprotein mainly responsible for scavenging cell-free hemoglobin (Hb). Although found in most bodily fluids of mammals, it is present in plasma at concentrations normally ranging from 0.5-3 mg/mL. After binding to cell-free Hb, the Hb-Hp complex is scavenged by CD163+ macrophages and monocytes to clear the organism of toxic cell-free Hb. Cell-free Hb toxicity is attributed to a variety of factors, including Hb extravasation into tissue space which elicits oxidative tissue injury, nitric oxide (NO) and peroxide scavenging, and free heme release. These factors can lead to acute and chronic vascular disease, inflammation, thrombosis, and renal damage. When bound to Hp, the large size of the Hb-Hp complex prevents Hb extravasation into the tissue space, lowering nitric oxide scavenging and vasoconstriction. Furthermore, Hp binding to Hb prevents heme release from Hb, and lowers Hb oxidative damage and inflammation. For these reason, Hp is used as a therapeutic in Japan and is being researched for treatment of various states of hemolysis. Recent studies have also shown various roles Hp plays as a chaperone and in regulation of redox states. In clinical settings, high levels of Hp are indicative of acute inflammation as its expression is upregulated in response to inflammatory cytokines, and low Hp levels are indicative of hemolysis due to receptor-mediated uptake of Hp-Hb complexes.
Hp is a polymorphic protein composed of αβ dimers in which the β polypeptide chain is coded by the same gene, while the α chain can be coded by either the Hp1 or Hp2 codominant alleles. These give rise to three main Hp phenotypes: Hp1-1, Hp2-1 and Hp2-2. The α and β chains are bound through disulfide bonds with the β, α-1 and α-2 chains having a molecular weight (MW) of 36, 9 and 18 kDa, respectively. The α-1 chain has a second cysteine residue after binding to the β chain that allows it to bind to another α chain of an αβ dimer. Thus, Hp1 homozygotes produce tetrameric Hp1-1 (two β and two α-1) species with a MW of about 89 kDa. On the other hand, the α-2 chain has two free cysteine residues when bound to the β chain allowing it to bind to two αβ dimers. This extra cysteine residue leads to the formation of Hp polymers in heterozygote or Hp2 homozygote individuals. Heterozygotes produce Hp2-1, a linear Hp polymer with an average MW of about 200 kDa, which can go up to about 500 kDa. Finally, Hp2 homozygotes produce Hp2-2 which is a cyclic polymer form of Hp with average MW of about 400 kDa ranging from 200 to 900 kDa. All types of Hp bind Hb via a practically irreversible reaction, with a Kd ranging from 1012 to 1015M. The bond occurs between the β chain of Hp to the β globin of Hb at a stoichiometry of 1:1 for each dimer. Since the MW of Hp differs between phenotypes, the mass stoichiometry is not consistent, with about a 1.3:1 mass binding ratio for Hp1-1:Hb and about a 1.6:1 mass binding ratio for Hp2-2:Hb. Furthermore, Hp 2-2 has been shown to have a higher affinity for the CD163+ receptor, but lower clearance rate through CD163+ uptake. However, the different Hp phenotypes reduce Hb toxicity to the same extent in vivo. Furthermore, Hp2-2 has not been found to have differences in the rate of heme loss, Hb oxidation, and Hb dimer association kinetics in vitro compared to Hpl-1 Although there are no significant differences with its role in Hb scavenging, differences in Hp phenotype have been associated with different rates of cardiovascular disease and cancer as well as different roles with some forms of disease.
In addition to the three main Hp phenotypes, another related Hb scavenging species is haptoglobin related protein (Hpr). Hpr is composed of smaller α and β chains than Hp1-1 and is predominantly found as single αβ dimers, but has been shown to form polymers. With >90% sequence identity to the Hp1 gene, Hpr binds to Hb with high affinity. Unlike Hp, the α chains do not covalently bind to other α chains through disulfide bonds to create αβ polymers, but are thought to connect via non-covalent interactions. The physiological role of Hpr differs from normal Hp as Hpr does not bind to the CD163 receptor and does not have increased expression during states of hemolysis. Instead, Hpr forms a complex with high-density lipoproteins called trypanosome lytic factor 1 (TLF1) and TLF2, which can have a large range of molecular sizes. These complexes have lytic activity against the African cattle parasite Trypanosoma brucei brucei, which use the trypanosoma receptor to obtain iron from the heme of Hb through binding of Hp-Hb complexes. Since Hpr can still bind to Hb in TFL1 and to the trypanosoma receptor, the Hpr-Hb in TFL1 acts as a “trojan horse” against trypanosoma parasites providing humans an innate defense against this disease.
The normal circulatory half-life of Hp is 1.5-2 days in humans, but the half-life of the Hp-Hb complex reduces to −20 min, with a maximum clearance rate of 0.13 mg/mL of plasma per hour. Due to the higher rate of clearance of the Hp-Hb complex, even though Hp production is upregulated during states of hemolysis, the concentration of Hp in plasma is inversely related to the concentration of cell-free Hb in plasma. In addition to its upregulation due to the presence of cell-free Hb in the circulation, Hp synthesis is heavily stimulated during acute phase reactions (inflammation, infection, trauma, and malignancy).
Hp can be used as a therapeutic during conditions that cause states of hemolysis (e.g., chronic anemia, transfusion, etc.). In these states, rupture of red blood cells (RBCs) releases cell-free Hb that can scavenge NO, leading to vasoconstriction as well as formation of free radicals and reactive oxygen species that can lead to oxidative damage of surrounding tissues. Hp upregulation during bacterial infection has been related to iron deprivation of pathogens. For this reason, Hp may be used to treat septic shock. Additionally, Hp or the Hp/serum protein mixtures presented here, may be used in RBC storage solutions to extend the ex vivo shelf-life of these cells by attenuating the side-effects associated with lysed RBCs. These solutions could also be co-administered with RBCs or hemoglobin-based oxygen carriers (HBOCs) to prevent and treat the side-effects associated with cell-free Hb.
Hp has been clinically approved in Japan since 1985. Reports of its use show positive effects against burn injuries, trauma from massive transfusions and as a prophylactic during surgical interventions such as cardiac bypass surgery with extracorporeal circulation. Hp treatment during severe burns has been shown to prevent acute renal failure and reduce kidney damage from surgery.
The biomedical applications of Hp are very promising. Its clinical use in Japan has shown positive effects against burn injuries, trauma from massive transfusions and as a prophylactic during surgical interventions such as cardiac bypass surgery with extracorporeal circulation. Hp treatment during severe burns has been shown to prevent acute renal failure, and reduce kidney damage from surgery. In general, Hp can be used to detoxify cell-free Hb that is present in the systemic circulation via various pathophysiological conditions or RBC transfusions that result in hemolysis. Hp treatment has also been shown to prevent damage from stored RBCs, potentially prolonging its shelf-life.
Yet, its wide-scale use is restricted by current production methods, which are not easily scalable and expensive. Furthermore, treatments with Hp require large quantities of material per dose. Existing Hp production protocols consist of using either Hb-affinity chromatography, hydrophobic interaction chromatography, anion-exchange chromatography, or recombinant Hp expression. Chromatography has low yields and is limited by the protein binding capacity of the column. Furthermore, chromatography requires the use of harsh denaturants to dissociate Hp from the bound chromatography matrix. Finally, recombinant Hp is expensive and cumbersome to manufacture.
This example describes a Hp production method via TFF that provides an easy, scalable and economically efficient method for producing Hp from plasma or plasma fractions. The plasma fraction chosen for this example was human Cohn Fraction IV obtained via the modified Cohn process of Kistler and Nitschmann (See, for example, Kistler, P. & Nitschmann, H. Large Scale Production of Human Plasma Fractions. Vox Sang. 7, 414-424 (1962)). This plasma fraction is known to contain large MW Hp (Hp2-2 and Hp2-1) from pooled plasma. Low MW Hp (small Hp2-1 polymers and Hp1-1) are primarily found in Cohn Fraction V.
Materials and Methods
Materials. Sodium phosphate dibasic, sodium phosphate monobasic, sodium chloride, and fumed silica (S5130) were purchased from Sigma Aldrich (St. Louis, Mo.). 0.2 μm Millex-GP PES syringe filters were purchased from Merck Millipore (Billerica, Mass.). A KrosFlo® Research II tangential flow filtration (TFF) system and hollow fiber (HF) filter modules were obtained from Spectrum Laboratories (Rancho Dominguez, Calif.). Human Fraction IV Paste was purchased from Seraplex, Inc (Pasadena, Calif.).
Hp Purification via TFF Without Fumed Silica. 500 g of human Fraction IV (FIV) paste from the modified Cohn process of Kistler and Nitschmann was suspended in 5 L of PBS, and homogenized in a blender. The resulting mixture was stirred overnight at 4° C. The ˜5 L solution was then centrifuged at 3700 g for 45 minutes to remove undissolved lipids. The supernatant was concentrated using a 0.2 μm hollow fiber (HF) filter to 2 L. The 2 L retentate was left to rest for 36 hrs to flocculate low density particles, while the filtrate was kept at 4° C. for further processing. After flocculation of the retentate, low density particles in solution were separated. The higher density fraction (Stage 0) was then concentrated to 800 mL on a 0.2 μm HF filter and subjected to 10 diafiltrations with PBS. The 0.2 μm filtrate (Stage 1) was concentrated to 150 mL and subjected to 100 diafiltrations on a 750 kDa HF filter using a mixture of the 0.2 μm permeate and PBS. The permeate of the 750 kDa (Stage 2) was then concentrated to 150 mL subjected to 40 diafiltrations using PBS on a 500 kDa HF filter. Finally, the permeate of the 500 kDa HF filter (Stage 3) was concentrated to 150 mL and subjected to 100 diafiltrations using PBS on a 100 kDa filter. A diagram of the purification process is shown in
Hp Purification via TFF with Fumed Silica. 500 g of human Fraction IV (FIV) paste from the modified Cohn process of Kistler and Nitschmann was suspended in 5 L of PBS, and homogenized in a blender. The resulting mixture was stirred overnight at 4° C. The ˜5 L solution was then centrifuged at 3700 g for 45 minutes to remove undissolved lipids. Fumed silica (Sigma Aldrich P #55130; St. Louis, Mo.) was then added to the sample at 20 mg/mL concentration and left stirring overnight at 4° C. The solution was then centrifuged to remove the silica agglomerates. Furthermore, the silica pellet was washed twice with PBS to maximize protein recovery. The fumed silica supernatant solution was then concentrated to 800 mL on a 0.2 μm HF filter and filtered for 15 diafiltrations. The 0.2 μm filtrate was concentrated to 150 mL and subjected to 100 diafiltrations using PBS on a 750 kDa HF filter. The permeate was then subjected to 40 diafiltrations using PBS on a 500 kDa HF filter. Finally, the permeate of the 500 kDa HF filter was subjected to 100 diafiltrations using PBS on a 100 kDa filter. A diagram of the purification process is shown in
Size Exclusion Chromatography: Samples were separated via size exclusion chromatography (SEC) using an Acclaim SEC-1000 (4.6×300 mm) column (Thermo Fisher Scientific, Waltham, Mass.) attached to a Dionex UltiMate 3000 system (Thermo Fisher Scientific, Waltham, Mass.). The mobile phase consisted of 50 mM potassium phosphate, pH 7.4 at a flow rate of 0.35 mL/min controlled by Chromeleon 7 software. Wavelength absorbance detection was set to λ=280 nm to detect protein, and λ=413 nm to detect Hb. To estimate the average MW of the Hp products, protein standards (conalbumin, 76 kDa; hHb, 64 kDa; carbonic anhydrase, 29 kDa; ribonuclease A, 14 kDa; and aprotinin, 6.5 kDa) were analyzed on the SEC column. The known molecular weight (MW) of the standards and their elution time were used to determine the coefficients (A, B) of a base 10 exponential function (MW=10A*(elution time)+B) via non-linear regression. The estimated function parameters were used to estimate the average MW of Hp products based on their elution time.
Hb Concentration. The concentration of Hb in the samples was measured spectrophotometrically via the Winterbourn equations.
Residual Hb in Hp Preparations. The residual Hb was quantified via the molar extinction coefficient of methemoglobin at its Soret Peak maxima of 404 nm (ε404=167 mM−1cm−1).
Residual Apohemoglobin in Hp Preparations. The concentration of residual apohemoglobin (apoHb) was estimated based on a modified version of the abridged dicyanohemin incorporation assay. Briefly, Hp samples were mixed with excess methemealbumin (heme bound to human serum albumin [HSA]) and the mixture left to react for 15 hours at room temperature. The change in absorbance of the reacted mixture compared to the initial sample components was used to estimate the amount of heme exchanged from methemealbumin to apoHb. Given the estimated extinction coefficients for the hemichrome-like apoHb species formed (ε412nm≈120 mM−1cm−1) and methemealbumin (ε412nm≈70 mM−1cm−1), the extinction coefficient for the change in absorbance spectra was determined to be 55 mM−1cm−1 at 412 nm.
ELISA. To quantify the concentration of residual protein components in the Hp samples, ELISA kits specific for Hp, transferrin (Tf), human serum albumin (HSA), and hemopexin (Hpx) were used (R&D Systems Catalog #DHAPGO for Hp, and Eagle BioSciences HTF31-K01 for Tf, HUA39-K01 for HSA, and HPX39-K01 for Hpx).
Gel Electrophoresis: The purity of Hp fractions was analyzed via SDS-PAGE using an Invitrogen Mini Gel Tank (Thermo Fisher Scientific, Waltham, Mass.). Wide-range Tris-Glycine gels consisting of pre-cast 4-20% or 10-20% polyacrylamide were used with samples prepared according to the manufacturer's guidelines. Gels were loaded with 20 μL of sample corresponding to approximately 30 μg of protein per lane and tested under reducing (via addition of 0.1 M DTT) and non-reducing conditions. Gels were stained for one and a half hours with Coomassie® Briliant Blue R-250 staining solution, then de-stained overnight. Gels were imaged on a table-top scanner at 300 dpi. To estimate the percent composition of each band, reduced and non-reduced gels were slightly overloaded (˜60 μg of protein) and densitometric analysis was performed on the scanned images using ImageJ. The Hp composition was determined based on the sum of protein bands corresponding to Hp or Hpr α and β chains from the reduced gels subtracted by the composition of the Hb-eluting band on the non-reduced gels (due to the known coelution of Hb α and β chains with the α-1 chain of Hp).
Trypsin Digest Mass Spectrometry. Samples were reduced with 5 mM DTT, incubated at 65° C. for 30 min. Iodoacetamide was then added to a final concentration of 15 mM and the sample was incubated in the dark, at room temperature for 30 minutes. Sequencing grade trypsin was added at a 1:50 ratio and the sample was digested overnight at 37° C. The following day, the samples were acidified with trifluoro acetic acid. The sample was clarified at 13,000 rpm for 5 min in a microcentrifuge, dried in a vacufuge and resuspended in 20 μL of 50 mM acetic acid. Peptide concentration was determined by NanoDrop (i.e. absorbance at 280 nm). Protein identification was performed using nano-liquid chromatography-nanospray tandem mass spectrometry (LC/MS/MS) on a Thermo Scientific Fusion Orbitrap mass spectrometer equipped with an EASY-Spray™ Sources operated in positive ion mode. Samples were separated on an easy spray nano column (Pepmap™ RSLC, C18 2μ 100 A, 75 μm×250 mm Thermo Scientific) using a 2D RSLC HPLC system from Thermo Scientific. Each sample was injected into the μ-Precolumn Cartridge (Thermo Scientific) and desalted with 0.1% formic acid in water for 5 minutes. The injector port was then switched to inject and the peptides were eluted off of the trap onto the column. Mobile phase A was 0.1% formic acid in water and acetonitrile (with 0.1% formic acid) was used as mobile phase B. Flow rate was set at 300 nL/min. Typically, mobile phase B was increased from 2% to 20% in 105 min and then increased from 20-32% in 20 min and again from 32%-95% in 1 min and then kept at 95% for another 4 min before being brought back quickly to 2% in 1 min. The column was equilibrated at 2% of mobile phase B (or 98% A) for 15 min before the next sample injection. MS/MS data was acquired with a spray voltage of 1.7 kV and a capillary temperature of 275° C. The scan sequence of the mass spectrometer was as follows: the analysis was programmed for a full scan recorded between m/z 375-1575 and a MS/MS scan to generate product ion spectra to determine amino acid sequence in consecutive scans starting from the most abundant peaks in the spectrum in the next 3 seconds. To achieve high mass accuracy MS determination, the full scan was performed at FT mode and the resolution was set at 120,000. The AGC Target ion number for FT full scan was set at 4×105 ions, and maximum ion injection time was set at 50 ms. MS/MS was performed using ion trap mode to ensure the highest signal intensity of MS/MS spectra using CID (for 2+ to 7+ charges). The AGC target ion number for ion trap scan was set at 1×104 ions, and maximum ion injection time was set at 30 ms. The CID fragmentation energy was set to 35%. Dynamic exclusion was enabled with a repeat count of 1 within 60 s and a low mass width and high mass width of 10 ppm. Data sets were analyzed as the Total Ion Intensity for each protein (normalized based on the total ion current) using Scaffold 4 (Proteome Software, Inc).
Total Protein Assay. Total protein of the samples was determined via the Bradford Assay.
Hb Binding Capacity of Hp (Fluorescence). The Hb binding capacity (HbBC) of samples was determined based on the fluorescence quenching method described in the literature. Briefly, a Hp sample is mixed with increasing amounts of Hb and the fluorescence emission at 330 nm (with excitation at 285 nm) is measured. Binding of Hb to Hp quenches the fluorescence of Hp, leading to an observable titration curve. This assay was repeated on products of Stage 2 and 3 for three different batches and compared to the SEC method to quantify HbBC.
Hb Binding Capacity of Hp (SEC): The difference in molecular weight (MW) between the Hp-Hb protein complex and pure Hb was used to assess the Hb binding capacity of Hp. Briefly, samples containing Hp were mixed with excess Hb then separated via SEC. The difference in the area under the curve between the pure Hb solution, and the mixture of Hb and Hp was used to assess the HbBC of Hp.
Heme Binding Activity: The activity of the heme-binding pocket of the protein scavenger cocktail was determined via the dicyanohemin (DCNh) incorporation assay. Briefly, increasing amounts of DCNh was added to a constant concentration of sample and the inflection point of the equilibrium absorbance at the Soret maxima was used to determine the molar quantity of heme required to saturate the heme-binding sites of the sample. The mass concentration of the heme-binding proteins was estimated based on an approximate molecular weight of 65 kDa for human serum albumin and hemopexin.
Effect of Fumed Silica: To assess the total loss of protein from the use of fumed silica, three 50 mL samples of suspended Fraction IV at 100 mg/mL were characterized after removal of the unsuspended FIV (mostly lipids), after addition of 20 mg/mL fumed silica with overnight stirring (fumed silica supernatant), and after each of the two washes (washing consisted of replacing the volume of supernatant removed with fresh PBS, and mixing the fumed silica pellet). The volume reduction due to fumed silica addition was approximated to be 15% as described by the manufacturer. Characterization consisted of quantification of HbBC and total protein, and via separation via SEC-HPLC. The percent of HbBC and total protein retained at each stage after fumed silica addition was calculated based on the ratio of the concentration of HbBC and total protein compared to the FIV supernatant prior to silica addition.
Results and Discussion
Hemoglobin Binding Capacity of Hp. Throughout this study, Hp activity was quantified based on the HbBC of the sample. This parameter indicates the mass of Hb that a given volume of the Hp sample can bind. This approach for Hp quantification is also used in Japan where the HbBC is equivalent to the international units (IU) of the therapeutic compositions of Hp. Given that different Hp phenotypes have different mass binding ratios to Hb and that the main role of Hp is as a Hb binding protein, the HbBC provides a more reliable and practical measurement of Hp activity in a Hp sample. This measurement of Hp activity is of critical importance in assessing the efficacy of various Hp production methods, especially those that rely on harsh denaturing conditions to purify Hp in which some of the Hp may be denatured.
There are various methods to quantify the HbBC of Hp samples. These include spectrophotometric titrations with Hb, immunodiffusion, gel electrophoresis, spectrophotometric differences between deoxygenated Hb and the deoxygenated Hb-Hp complex, differences in peroxidase activity of Hb compared to the Hb-Hp complex, and fluorescence titration of Hp with Hb. Unfortunately, spectrophotometric assays can be dependent on the Hp phenotype and convoluted by other species. Additionally, ELISA can also be used, but differences in polymer sizes can also lead to inaccurate readings. One of the initial methods was based on the fluorescence quenching of Hp tryptophan residues upon Hb binding. In this method, a stock Hp sample is titrated with increasing Hb concentrations and the fluorescence intensity is measured after each addition. The saturation point of Hb binding sites in Hp is determined by the change in slope of the titration curve.
Comparing the fluorescence titration and HPLC-SEC AUC Hp quantification methods for six different Hp containing samples, there was less than 5% variation of the results which, at the concentrations tested, led to less than 1 mg/mL of HbBC variation. Furthermore, the HPLC-SEC method has less than 1% variation in precision. Thus, the 5% variation seen may have been due to the intrinsic variation of the fluorescence titration method. Furthermore, these variations were similar or better than the reported values for previous HbBC quantification methods. The difference in peroxidase activity, had larger variations (7.6% using different standards and 2.6% using same standard curve). Spectrophotometric differences between free Hb and Hb-Hp had more than 10% error. Furthermore, spectrophotometric titration with Hb had about 2% variation within the same sample and ranged from 2-11% when the same sample was tested on a different day. A similar method which employed SEC-HPLC to determine HbBC has been employed previously, in which the AUC of the Hb-Hp complex was divided by the total AUC of the chromatogram. Although relying on the same concept (use of HPLC-SEC to separate Hb from the Hp-Hb complex), the previously used method requires that the Hp does not have any Hb bound to it. Furthermore, slight modifications in the absorbance of Hb-bound species may occur. For example, the change in absorbance from the binding of free cyanomethemoglobin to Hp has been used to quantify HbBC. Using the method presented here in which only the AUC of the Hb peak is used in the analysis, this method removes potential errors from analysis of the AUC from the Hp-Hb complex peak.
TFF Production of Hp Without Fumed Silica. Production of Hp mixtures via the tangential flow filtration (TFF) procedure described in the Methods Section yielded 5 different stages in which different molecular weight (MW) proteins could be isolated. These 5 stages were the proteins retained on the 0.2 μm HF filter (Stage 0, 0.2 μm retentate), the bracket of proteins between the 0.2 μm and 750 kDa HF filters (Stage 1, 0.2 μm-750 kDa), the bracket between the 750 kDa and the 500 kDa (Stage 2, 750-500 kDa), the bracket between the 500 and 100 kDa HF filters (Stage 3, 500-100 kDa), and lastly, the permeate of the 100 kDa HF filter (Stage 4, <100 kDa). The protein and Hb binding capacity (HbBC) yield at each stage based on the average of four batches without the use of fumed silica is shown in Table 4. The recovery data for total protein and Hb binding capacity (HbBC) at each stage of processing is also shown in
From the results of Table 4 and
Given that most serum proteins have MW smaller than 100 kDa, most of the suspended protein permeated through the 100 kDa filter, leading to 65% of the soluble protein being lost in Stage 4. Furthermore, since Hp can be present over a wide MW range and that FIV contains mostly polymeric Hp, Stage 3 retained a high HbBC fraction of FIV, indicating that Hp was present in this >100 kDa MW bracket. Yet, given that only a small fraction of the total protein of FIV was retained in Stage 3, this stage had the highest HbBC per milligram of total protein. This high HbBC to total protein ratio indicated a high Hp purity in Stage 3. Although at similar HbBC to total protein ratio, only a small quantity (˜4 mL at ˜50 mg/mL) of Stage 2 was purified, with most of the product begin retained in Stage 3 (˜38 mL at 100 mg/mL). Low retention in Stage 2 suggested that the Hp polymers primarily permeated through the 500 kDa filter. Yet, due to the separation on a 500 kDa filter, Stage 2 contained mostly high MW (HMW) Hp polymers, while Stage 3 contained mostly lower MW (LMW) Hp which may allow for testing of the effect of Hp polymer size on the therapeutic index in various disease states. Based on these MW ranges, Hp polymers in Stages 2 and 3 mainly consisted of a mixture of Hp2-2 and Hp2-1, since any Hp1-1 present in FIV (expected to be primarily present in Cohn Fraction V) was too small to be retained in Stage 3.
Unfortunately for the purification of Hp, not only did most proteins permeate through the 100 kDa system, but almost 50% of the initial HbBC also permeated through the system, indicating that these Hp species in FIV were also permeable through the 100 kDa filter (small Hp polymers). Therefore, decreasing the numbers of diafiltrations at the last stage or using a smaller final MW filter could improve retention of these species with a potential drawback of reducing product purity. Furthermore, a large fraction of the initial HbBC (28%) and total protein (29%) was retained on the 0.2 μm filter, indicating that this filter was likely fouled during processing. Thus, starting with a different form of processed human plasma or removing some of the initial fouling particulates may benefit the initial filtration step. Possible solutions could be to introduce a larger filter pore size prior to the 0.2 μm filter or precipitating the protein suspension using ammonium sulfate or other salting-out agents. One promising fraction to use as the starting material could be Cohn Fraction IV-4, which removes many of the lipoproteins from the starting material. Furthermore, as shown later in this study, addition of fumed silica greatly enhanced filtration through the 0.2 μm stage. Stage 1 (0.2 μm-750 kDa) also showed potential as a product, but the sample may have to undergo further PBS diafiltration. Furthermore, via this method, Stage 1 contained large molecular weight proteins that did not seem to bind any Hb (see HPLC-SEC).
HPLC-SEC was performed on each of the processing stages and the results are shown in
In
From these curves, it was also possible to note that some of the high-MW protein species (elution times of ˜7.5 min or earlier) in Stages 0, 1, 2 and 3 did not show Hb-binding properties. This was noted by the lack of increase in absorbance at 280 nm of these high-MW species when excess Hb was added to the sample. When Hb binds to Hp forming Hb-Hp complexes, the absorbance at the Hp/Hp-Hb elution time increases due to the absorbance of Hb at 280 nm. Thus, if no increase was observed, it indicated that most of these high-MW species did not bind to Hb and were likely impurities that were not removed during the TFF processing. Interestingly, Stage 1 (0.2 μm-750 kDa) may also have therapeutic potential, but the sample had large quantities of large non-Hb binding proteins and may have to undergo further PBS diafiltrations, increasing overall processing time.
To analyze the protein species in each of the processing stages, SDS-PAGE was performed under both reducing and non-reducing conditions. Furthermore, protein identity was confirmed via trypsin digestion and mass spectroscopy (MS) of the samples. The gels from one representative batch and the percent composition based on the label-free quantitative MS analysis is shown in
From the results of
Based on the increase in relative intensity of the polymeric species in Stages 1-3 of the non-reduced SDS-PAGEs, it was noted that the TFF process was capable of purifying high-MW species present in the FIV suspension (Stage 0). Although these species likely consisted of mostly Hp polymers, other large MW protein species and/or polymerized proteins may also have contributed to the “Polymers” band of
Hemoglobin (Hb) and apolipoproteins were also present under the non-reducing SDS-PAGE shown in
On the reduced SDS-PAGE, Hp dissociated into its α and β components. As expected, the majority of the α chains were α-2 which allow for the polymerization of Hp. Furthermore, a higher intensity of the band at ˜60 kDa was detected, likely due to reduction of disulfide bonded albumin polymers and full dissociation of proteins associated with lipoproteins. The presence of these polymeric albumin species would explain the detection of 4-5% HSA via MS on Stages 2 and 3 (
Protein purity was also estimated via densitometric analysis of the bands of Stage 2 (HMW) and 3 (LMW). The results are shown in Table 5.
From the densitometry assessment, Stage 2 and 3 were composed of ˜70 and ˜75% Hp, respectively. The major impurity were proteins in the 55-65 kDa range with ˜20 and ˜10% in Stages 2 and 3, respectively. Based on MS, these impurities were primarily composed of human serum albumin (HSA) and α-1 antitrypsin (AT). Due to their similar molecular weights, the proteins were not separately quantified via densitometry. But MS indicated that Stage 2 and 3 had 19% and 11% of AT with 4% and 5% of HSA, respectively. Unfortunately, many impurities were present in the Hp samples, which may have contributed to an inaccurate estimate of Hp purity. Most of these impurities likely originated from proteins associated with lipoproteins and/or proteins that can undergo polymerization. Interestingly, these impurities showed little deviations from batch-to-batch, demonstrating that the purification process was reproducible.
Although convoluted from contaminant proteins, the Hp purity from SDS-PAGE was similar to the expected Hp content based on the average HbBC per mg of total protein of Stage 2 and 3. Using the theoretical the mass ratio of Hp2-2 to Hb, the expected purity of the samples would be approximately 50% and 60% for Stages 2 and 3, respectively. Yet, the Bradford assay may have led to overestimation of total protein due to high concentration of glycoproteins which can also react with the dye used in the assay (Hp can have ˜20% of its total mass attributed to conjugated carbohydrates). Moreover, high MW Hp polymers isolated from serum have been shown to have even higher mass binding ratios than theoretical (>2:1 Hb:Hp), potentially due to tertiary structure steric hindrance. At a mass binding ratio of 2, Stage 3 would have ˜70% Hp, similar to what was obtained from densitometric analysis.
From UV-visible spectrometry, residual Hb contributed ˜1% of the protein mass for the LMW and HMW stages. Yet, based on the SDS-PAGE densitometry, Hb chains consisted of ˜3% of the total mass, indicating that some of the Hp may be bound to apohemoglobin (apoHb). Thus, the residual apoHb content was assessed by adding excess heme-albumin and monitoring the increase in absorbance at the Soret maxima as described in the Methods section. Given that Hp has been shown to not bind appreciably to heme, the change in absorbance was due to heme exchange from HSA to apoHb. Heme binding indicated that some of the residual Hb may have had its heme extracted during the acidic-ethanol fractioning of plasma to obtain Fraction IV. This heme-binding property of the purified Hp sample could be beneficial during hemolytic states in which free heme is also present.
TFF Purification of Hp with Fumed Silica. From the results already shown, it was apparent that lipoproteins were likely the carriers for a large proportion of the impurities in the sample. Fumed silica is commonly used for de-lipidation of serum samples as it is capable of adsorbing lipids. Thus, the effect of using fumed silica on FIV was assessed, and the results are shown in
The analysis of percent recovery (
From the results of Table 6 and
The HPLC-SEC chromatograms followed the same trends for Stages 2 to 4 as the batches that did not use fumed silica. More importantly, Stage 2 and Stage 3 had a reduced left tail-end of the non-Hb-binding proteins but the estimated molecular weight of Stages 2 and 3 increased compared to without the use of fumed silica. Stage 2 and Stage 3 had an average MW of 520±30 kDa and 390±20 kDa, respectively. Thus, even though there was removal of the large ˜7.5 min eluting species, the flux of large MW proteins through the filters improved, yielding larger average sized proteins in each stage. The increase in protein permeation through the filters is also shown by the lack of a peak at ˜9.2 min on Stage 0, indicating high passage of these ˜100 kDa species. Yet, the presence of low MW protein species in Stage 0 indicated that the filter was still fouling. The samples of the main Hp stages (Stage 2 and 3) were also analyzed via SDS-PAGE and trypsin digest MS, the result is shown in
From the SDS-PAGE, it was apparent that these fractions were primarily composed of Hp. Little of the impurities from the purification method without fumed silica were detected. From a low protein loaded gel (shown in
From the purity assessment in Table 7, the BMW fraction was composed of ˜80% pure Hp and the LMW fraction was composed of ˜90% pure Hp. Thus, in agreement with the higher HbBC per mg, it was confirmed that the purity of the two main Hp products had greatly improved with the use of fumed silica. Yet, the purity of Hp is likely higher due to the slight overloading required to detect these impurities on the SDS-PAGE. Furthermore, similar to the process without the use of fumed silica, the method yielded consistent product compositions as demonstrated by the small deviations from densitometric analysis.
Interestingly, based on the average HbBC to total protein ratio and the Hb mass binding ratio of Hp2-2, the expected purity of the LMW Hp fraction (Stage 2) would be ˜85%. Unlike the process without the use of fumed silica, this value was in close agreement with the SDS-PAGE densitometry, indicating that very few protein contaminants were present in the samples. Furthermore, as explained before, the mass binding ratio between Hp2-2 and Hb may be larger than the theoretical value, leading to a closer Hp purity of >95% in agreement with densitometric analysis of SDS-PAGE.
Moreover, the higher purity from the Hp product purified with the use of fumed silica indicated that either the polymerized forms of the non-Hp proteins have a high affinity for the silica (potentially due to higher hydrophobicity from partial protein unfolding) or that most of these proteins were associated with the lipoproteins. Practically, no change in the α-2 macroglobulin fraction was observed indicating that this multimeric (720 kDa) protein species did not have a high affinity for the silica particles.
ELISA was performed on Stages 2 and 3 of the fumed silica treated batches to determine the mass percentage composition of Hp, HSA, and Tf to compare to the values determined by densitometric analysis of the SDS-PAGE. Unfortunately, the Hp ELISA was unable to accurately quantify the large MW Hp polymers in Stages 2 and Stage 3. The ELISA kit resulted in a Hp mass composition of 21±0.2% and 34±0.2% for Stages 2 and 3, respectively. As discussed previously, Hp ELISAs may not provide proper quantification of large MW Hp species. These large MW species may have sterically inhibited binding of Hp to immobilized Hp antibodies. The HSA content was determined to be 0.8±0.01% and 0.8±0.2% for Stages 2 and 3, respectively. This result agreed with MS data that demonstrated relatively similar HSA content for Stages 2 and 3. Thus, the difference in composition for the HSA/AT band on the reduced SDS-PAGE was mainly attributed to the higher AT composition in Stage 2 as seen by the high AT total ion intensity in the MS. The Tf composition was determined to be 0.5±0.09% and 0.3±0.09% for Stages 2 and 3, respectively. These results were similar to the values determined by SDS-PAGE, which showed <1% of Tf for both Stage 2 and Stage 3. Finally, ELISA kits were used to confirm that the hemichrome species formed from heme exchange from heme-HSA to the Hp sample was due to residual apoHb and not residual Hpx. Although Hpx was not detected on MS or apparent on the SDS-PAGE, both denatured apoHb and Hpx form similar hemichrome spectra (bis-histidine bonded heme). Hence ELISA results showed that the Hpx content was 0.1±0.02% and 0.1±0.04% of the total protein for Stages 2 and 3, respectively. Given that the calculated apoHb mass content was ˜3%, practically all heme exchange was due to residual apoHb in Stages 2 and 3 with a minor contribution from Hpx. Furthermore, since Hpx has a larger MW (on a heme basis) than apoHb, if Hpx was the heme-binding species, a mass content of >10% Hpx would have been determined
Although the purified samples were not composed of only Hp, the extra proteins in the HMW and LMW fractions yield a protein cocktail potentially useful for treatments of hemolysis. For example, the α-2 macroglobulin (α2M) protein is a broad specificity protease inhibitor. Furthermore, α2M helps maintain a balanced clotting system by both inhibiting the coagulating protein thrombin and inhibiting the anti-coagulating Protein C system. These characteristics may improve the effectiveness of the Hp product for applications in which the patient has an abnormal balance of the clotting proteins. Furthermore, α2M can help maintain hemodynamic equilibrium after scalding/burning by inhibiting prostaglandin E2 (vasodilator) and restricting loss of plasma volume. Moreover, the anti-inflammatory, anti-fibrosis and anti-oxidant functions of α2M have been linked to its role as a radioprotective agent. These characteristics can improve treatment of hemolytic states due to burn injury or radiation injury. Finally, α2M along with AT have been shown to mediate the binding of Tf to its surface receptor. In doing so, α2M can help with the removal of excess iron potentially released during prolonged states of hemolysis.
For the Hp samples purified without fumed silica, the presence of HSA, Tf and AT may improve the therapeutic effect of the protein mixture compared to pure Hp. HSA is a multifunctional protein, with major roles in the regulation of acid-base balance, oncotic pressure, binding/transport of endogenous and exogenous molecules and drugs (binds to heme upon depletion of hemopexin, potentially improving hemolysis treatment with the Hp protein cocktail), protection against exogenous toxins, maintenance of microvascular integrity and capillary permeability, antioxidant and anticoagulant activity. AT and transferrin have also been shown to bind to heme, which is a highly oxidative by-product of Hb degradation during states of hemolysis. AT, named for its ability to inhibit trypsin, can also inhibit other proteases, which is also known as: α1-antiprotease inhibitor. Due to its anti-proteolytic function, AT has general anti-inflammatory properties. In addition, AT plays a large role in vivo by preventing lung damage via inhibition of neutrophil elastase. Co-treatment of Hp with AT may constitute an improved treatment of pulmonary hypertension as both Hb (and its by-products) and neutrophil elastase have been shown to have deleterious effects. Finally, Tf is an antioxidant protein responsible for iron binding and transport. Thus, iron build-up due to excessive hemolysis could be neutralized by Tf.
As stated previously, apolipoprotein A-1 is the main component of high-density lipoproteins (HDL) and is found in the TLF1 complex (contains Hpr that binds to Hb). HDL therapy has been shown to treat atherosclerosis, improving blood flow and the HDL (expected to be the form in the protein cocktail on the process without fumed silica) has had greater clinical efficacy than pure apolipoprotein A-1 (likely due to its short half-life). In addition to its well-known role in lipid transport, HDL/apolipoprotein A-1 has various pleiotropic effects such as antioxidant, anti-inflammatory, antithrombotic (anticoagulant and increased NO bioavailability) and vasoprotective activities. Furthermore, HDL has been shown to negate the effects of lipopolysaccharides, reducing its pro-inflammatory responses. Finally, apolipoprotein A1 has also been shown to have antimicrobial activity.
To reduce the costs associated with the use of buffers, future studies may aim to optimize the number of diafiltrations at each stage for effective protein transmission and protein purity. Furthermore, by retaining the proteins in Stage 4 using a low MW TFF filter (<50 kDa), the permeated buffer may be used for future processing as it would contain little to no proteins. Bacterial contamination of the buffer is minimized given the filtration through the 0.2 μM and 750 kDa HF filters at the start of the next batch. The fraction retained in Stage 4 may also have therapeutic uses as it provides a promising mixture or proteins. Moreover, the protein mixture in Stage 4 could be used as the starting material for conventional chromatography purification to yield low MW Hp. Finally, the initial protein loading of FIV can also be increased. Dissolving 1 kg of FIV into 5 L of PBS before processing can yield approximately double the Hp yield with no discernable difference in purity compared to that of the 500 g batches.
Although the starting material for FIV consisted of pooled human plasma, which may have safety risks associated with infectious agents, the combination of Cohn acid-ethanol fractionation and TFF processing inherently reduces these risks. TFF clarification with the 0.2 μm and 750 kDa HF filters remove most pathogenic bacteria. In the case of viruses, the Cohn acid-ethanol fractionation process provides an approximate 4 log10 reduction value (LRV) for various viruses. Furthermore, nanofiltration using TFF can add an additional 5 (LRV). Finally, the Hp sample may undergo a final virus reduction step if desired such as solvent/detergent or pasteurization to reach the desired level of pathogen reduction.
One drawback for scalability of the current process is the use of centrifuges for removal of undissolved lipids and/or fumed silica. Cloth or depth filtration may substitute for the centrifugation step used to remove the unsuspended lipids. Furthermore, continuous centrifugation may be employed to separate the fumed silica, as it requires low relative centrifugal force to form a pellet. Finally, given that in both centrifugation steps, the goal was to remove lipids, these steps could be combined into a single centrifugation step to decrease processing time.
Conclusion
Overall, starting from 500 g of Cohn Fraction IV paste and without the use of fumed silica, 1.2 g of HbBC at ˜75% purity was obtained in solutions with ˜100 mg/mL total protein and 33 mg/mL HbBC. With the use of fumed silica, the yield increased to 1.7 g of HbBC at >95% purity in solutions with ˜100 mg/mL total protein and ˜52 mg/mL HbBC.
Future studies may aim at improving processing time by optimizing the number of diafiltrations and buffer usage. Taken together, this study presents a novel and improved method for producing large quantities of large MW Hp (mixture of Hp2-2 and Hp2-1) at >95% purity
This example describes a process to purify a target protein (TP) from a mixture of proteins by exploiting molecular size changes that arise from the formation of a protein complex consisting of the TP and the TP binding molecule (TPBM). Briefly, the method employs tangential flow filtration (TFF) with a defined molecular weight cut off (MWCO) membrane to first permeate the TP+other protein impurities (filtrate) that are below the MWCO of the membrane, as well as set the maximum size/molecular weight of the protein species in the filtrate. A TPBM (could be another protein, antibody, aptamer or some other molecule) is then added to the filtrate to selectively create a protein complex with the TP in the protein mixture that is above the MWCO of the original membrane. With only the complexed TP above the MWCO of the original membrane, it can be selectively separated from the other low MW protein components of the filtrate using the original MWCO membrane. TFF can then be applied to buffer exchange the complexed protein under dissociative conditions and separate the TP from the TPBM using a MWCO membrane that is between the MW of the TP and TPBM.
This theoretical strategy is schematically illustrated in
General Strategies
Example Strategy 1—Purification of a 20 kDa TP using IgG antibody specific to TP. In a mixture of cell lysate (may need prior clarification through 0.2 micron filter and/or 50 nm filter), filter all material through a 70 kDa MWCO membrane. Add immunoglobulin G (IgG, commonly used antibody type) antibody specific to the TP into the filtrate. This will create an antibody-TP protein complex with MW >70 kDa (˜190 kDa, assuming two antigen binding-sites per antibody). The ˜190 kDa protein complex with the 20 kDa TG is now in a mixture with other proteins <70 kDa. Thus, this solution can be re-filtered through the 70 kDa MWCO membrane to retain the TP-antibody protein complex. The isolated TP-antibody complex can then be buffer exchanged into appropriate conditions to dissociate the TP-antibody complex (i.e. altered pH, salt concentration, or other appropriate denaturing condition). With the TP dissociated in solution from the antibody, refiltering the solution through the 70 kDa MWCO membrane will lead to the 20 kDa TP in the filtrate, and the 150 kDa antibody will be in the retentate. With the TP and antibody isolated, these species can be buffer exchanged into appropriate buffers on 10 kDa and 70 kDa membranes, respectively to yield purified TP and antibody. It is important to note that polyclonal antibodies may form aggregates at equimolar concentrations, thus excess target protein or excess antibody can be used for purification with these antibody species. On the other hand, monoclonal antibodies would not have the same issue as they only form specific complexes.
Example Strategy 2—Purification of a 200 kDa TP Using IgG (Non-Reduceable Protein, IgM or Equivalent Large TPBM could be Used for its Purification as in Example Strategy 1) Antibody Specific to TP. In a mixture of cell lysate (may need prior clarification through 0.2 micron filter and/or 50 nm filter), filter all material through a 300 kDa MWCO membrane. Add IgG antibody specific to TP of interest into the filtrate. This will create a TP-antibody protein complex with MW>300 kDa (˜350-670 kDa). The 200 kDa TP is now in an >300 kDa TP-antibody protein complex in a mixture with other proteins <300 kDa. Re-filter the protein solution through a 300 kDa MWCO membrane. The TP-antibody complex will be retained on the 300 kDa MWCO membrane. If the TP does not dissociate from the TP-antibody complex under reducing conditions, reduction of IgG will allow for its separation from the TP on a 100 kDa filter. The 200 kDa TP will be retained on the 100 kDa MWCO membrane, while the reduced components of IgG will enter the permeate. Both the TP and IgG components can be diafiltered into appropriate buffers. It is important to note that polyclonal antibodies may form aggregates at equimolar concentrations, thus excess target protein or excess antibody can be used for purification with these antibody species. On the other hand, monoclonal antibodies would not have the same issue as they only form specific complexes.
Example Strategy 3—Tagged recombinant proteins. Recombinant proteins can be synthesized with tags that facilitate their purification outlined in the Example Strategies above, removing the requirement for affinity columns. Instead of binding to affinity beads in columns, tags would bind to molecules that have larger MW than the MWCO of the employed membrane. For example, in a mixture of cell lysate with the strep-tagged 10 kDa recombinant protein (TrP), filter all material through a 30 kDa MWCO membrane. Add streptavidin to create a TrP-streptavidin complex with MW>60 kDa. The ˜60 kDa TrP-streptavidin complex is now in a mixture with other proteins <30 kDa. Thus, this solution can be re-filtered through the 30 kDa MWCO membrane to retain the TrP-streptavidin complex. The isolated protein complex can be buffer exchange into appropriate conditions to dissociate the protein complex. With the TrP dissociated in solution from the streptavidin, refiltering through the 30 kDa MWCO membrane will lead to the 10 kDa TrP in the filtrate, and the 50 kDa streptavidin will be in the retentate. With the TrP and streptavidin isolated, these species can be buffer exchanged into appropriate buffers on 1 kDa and 30 kDa membranes, respectively to yield purified TrP and streptavidin. This strategy employs currently known tagged recombinant protein technology, but improvement of the technologies for the proposed TFF purification could greatly ease its use. For example, polymerizing or developing large maltose/streptavidin/glutathione-bound molecules could improve applicability of this size exclusion method with currently used tagging systems (e.g., maltose-binding protein, strep-tag, glutathione-S-transferase, split-intein, etc.).
Purification of Haptoglobin (Hp) from Human Plasma or a Plasma-Derived Fractions Via Protein Complex Formation.
The theoretical manufacturing scheme for the presented technology is shown in
500 g of human Fraction IV (FIV) paste from the modified Cohn process of Kistler and Nitschmann was suspended in 5 L of PBS, and homogenized in a blender. The resulting mixture was stirred overnight at 4° C. The ˜5 L solution was centrifuged for 40 min at 3700 g to remove insoluble particulates (mostly lipoproteins). Then, the supernatant was concentrated using a 0.2 μm hollow fiber filter to 2 L The retentate was left to rest for 36 hrs to flocculate low density particles, while the filtrate was kept at 4° C. for further processing. After flocculation of the retentate, low density particles in solution were separated. The higher density fraction was then concentrated ˜1 L on a 0.2 μm hollow fiber filter and subjected to 10 diafiltrations with PBS. The 0.2 μm filtrate was concentrated to 150 mL and subjected to 100 diafiltrations on a 750 kDa hollow fiber filter using a mixture of the 0.2 μm permeate and PBS. The permeate was then subjected to 40 diafiltrations using PBS on a 500 kDa hollow fiber filter. Finally, the permeate of the 500 kDa hollow fiber filter was subjected to 100 diafiltrations using PBS on a 100 kDa hollow fiber filter (intermediate stages comprising of 750 and 500 kDa hollow fiber filtration were employed prior to the 100 kDa hollow fiber filter to avoid filter fouling). Hemoglobin (Hb) was then continuously added to the permeate from the 100 kDa hollow fiber filter to form the Hb-Hp complex, while maintaining the solution with excess Hb to bind all Hp in the permeate. The filtrate/Hb mixture was then subjected to diafiltration (100 or 200×) on a 100 kDa hollow fiber filter using fresh PBS to remove excess Hb and low molecular weight (MW) proteins. The resulting Hb-Hp complex was then centrifuged for 30 min at 3000 g to remove any insoluble particulates. The 100 diafiltration trial yielded 200 mL of 2 mg/mL Hb-Hp complex, while the 200 diafiltration trial yielded 200 mL of 0.8 mg/mL Hb-Hp complex. The diagram for the purification process is shown in
To facilitate dissociation of Hb from the purified Hb-Hp complex, 7 mL of Hb-Hp at 2 mg/mL was buffer exchanged (7 diacycles) into a 5 M Urea solution at a pH 10 using a 70 kDa hollow fiber filter. The resulting unfolded protein mixture was then subjected to 10×diafiltration using the urea solution with a rest period of 12 hr in between processing to yield a total of 30 diacycles. The solution was then diafiltered for 10 diacycles into DI water followed by 7 diacycles into PBS using a 30 kDa hollow fiber filter. The schematic of this process is shown in
The SDS-PAGE of the purified Hb-Hp complex and recovered pure Hp from one batch is shown in
From the SDS-PAGE, practically no impurities could be detected (>98% pure) for the purified Hp-Hb. Furthermore, from both SDS-PAGE band intensity analysis and the spectrophotometrically determined amount of Hb bound to the purified Hp species, the Hp to Hb mass binding ratio was calculated to be 1.6:1. This is the same mass binding ratio assuming one Hp2-2 dimer is bound to one Hb dimer. Urea treatment was not successful in removing all of the bound Hb. Using the 1.6:1 mass binding ratio, SDS-PAGE analysis indicated that about 20% of the Hp was still bound to Hb. In comparison, using total protein and spectrophotometric analysis, the product consisted of 25% active Hp, 29% Hb-Hp complex and 52% inactive Hp (denatured). Furthermore, compared to the starting Hb-Hp complex, 52% of Hp was lost during diafiltration, 12% was active, 13% remained bound to Hb and 23% was denatured. These results could be improved through optimization of the protein unfolding conditions in urea to avoid protein denaturing and via selection of a lower MWCO hollow fiber filter for the diafiltrations to avoid loss of protein.
From the Hb binding fluorescence assay, the final solution had a Hb binding capacity of 0.5 mg/mL, which indicated that only 10% of the initial Hb binding capacity was recovered. This large loss of Hp was attributed to loss of protein and protein denaturing during urea diafiltration as well as retained Hb in the product.
During the purification of the Hp-Hb complex, samples were taken at different stages of the process. These samples were analyzed on an HPLC-SEC column and the results were compared to the theoretically predicted separation based on the schematic in
From these results, the addition of Hb to Fraction IV increased the amount of large molecular weight species (compare 1 to 1*). These species matched our purified product (4). Furthermore, the permeate analysis (2) showed that the unbound Hp did not easily permeate the 100 kDa hollow fiber filter. This can be seen due to a lower relative abundance of the Hb-Hp complex when Hb was added comparing Fraction IV to the permeate (1* to 3). Another observation was that, by comparing 2 to 4, it was noticeable that the Hb-Hp complex was capable of permeating through the 100 kDa hollow fiber filter. This observation agreed with the different Hb-Hp yields obtained via 100× or 200 diafiltrations. Combining the chromatograms into one figure better represented the statements regarding retention of unbound target Hp and permeation of the target Hb-Hp complex. The combined elution chromatograms at 280 and 413 nm wavelength detection is shown in
The general mechanism of action for the ability of the apoHb-Hp complex to scavenge cell-free Hb and heme is shown in
Apohemoglobin-Haptoglobin Complex Preparation. The apoHb-Hp complex can be made by reacting apoHb with Hp. The high binding affinity drives the reaction for complex formation. A Hp solution with a Hb binding capacity (HbBC) of 24 mg/mL was mixed with an apoHb solution with 37 mg/mL of active apoHb at a 1:1 and 1:4 volume ratio. The resultant mixture was separated on a size exclusion chromatography (SEC) column for analysis. Large molecular weight Hp (Hp2-2 and Hp2-1) was mixed with apoHb with a molecular weight of 31 kDa (dimeric apoHb) and separated on an analytical Acclaim SEC-1000 (4.6×300 mm) column (Thermo Fisher Scientific, Waltham, Mass.) attached to a Dionex UltiMate 3000 system (Thermo Fisher Scientific, Waltham, Mass.). The mobile phase consisted of 50 mM potassium phosphate, pH 7.4. The flow rate and UV-visible spectral detection was controlled using Chromeleon 7 software with detection set to A=280 nm to detect protein elution at a flow rate 0.35 mL/min. The percent change of the area under the curve between a pure apoHb solution and a mixture of apoHb-Hp with excess apoHb was used to determine the percentage of apoHb that was bound to Hp. This percentage was compared to the mass of pure apoHb loaded to determine the Hp binding capacity of apoHb. This value was compared to the HbBC of the Hp sample.
Results
As seen in
The apoHb-Hp complex can be made to have pure complex or excess of one of the species (apoHb or Hp). Excess of either apoHb or Hp may allow for targeted treatment of different conditions characterized by higher free heme or free Hb.
Haptoglobin (Hp) is the plasma protein that binds and clears cell-free hemoglobin (Hb), while apohemoglobin (apoHb, i.e. Hb devoid of heme) can bind heme. Therefore, the apoHb-Hp protein complex should facilitate holoHb-apoHb dimer exchange and apoHb-heme intercalation. Thus, it was hypothesized that apoHb-Hp could facilitate both Hb and heme clearance, which if not alleviated could have severe microcirculatory consequences. In this example, apoHb-Hp and Hb/heme ligand interactions were characterized, and their in vivo consequences were assessed. Hb exchange and heme binding with the apoHb-Hp complex was studied with transfer assays using size exclusion-high performance liquid chromatography coupled with UV-visible spectrophotometry. Exchange/transfer experiments were conducted in guinea pigs dosed with Hb or heme-albumin followed by a challenge with equimolar amounts of apoHb-Hp. Finally, systemic and microcirculatory parameters were studied in hamsters instrumented with a dorsal window chamber via intravital microscopy. In vitro and in vivo Hb exchange and heme transfer experiments demonstrated proof-of-concept Hb/heme ligand transfer to apoHb-Hp. Dosing with the apoHb-Hp complex reversed Hb- and heme-induced systemic hypertension and microvascular vasoconstriction, reduced microvascular blood flow and diminished functional capillary density. Therefore, this example highlights the apoHb-Hp complex as a novel therapeutic strategy to attenuate the adverse systemic and microvascular responses to intravascular Hb and heme exposure.
Introduction
Erythropoiesis and hemolysis occur at similar rates in healthy organisms, thus maintaining the total population of red blood cells (RBCs) in the circulation. RBC homeostasis results in the balanced cycle of erythropoiesis and hemolysis, along with the effective transport and clearance of hemoglobin (Hb) and its degradation products (heme and iron). Genetic hemolytic diseases (e.g. sickle cell anemia and thalassemia), acquired hemolytic infections (e.g. gram positive and malarial hemolysins) as well as hemolytic xenobiotic toxins (e.g. heavy metals, dapsone and phenyl hydrazine) damage the RBC, disrupting this delicate equilibrium by increasing the rate of hemolysis. Increased intravascular hemolysis and the Hb degradation product heme progress hemolytic diseases, such as sickle cell anemia, malaria and sepsis, which affect millions of patients every year.
The presence of Hb outside of the erythrocyte's protective physiologic system exposes Hb to a structurally unstable environment due to the increased dilutional dynamic equilibrium between tetrameric (α2β2) and dimeric components (αβ), which are more prone to autoxidation and extravasation relative to tetrameric Hb. Acellular Hb (α2β2) dissociation into αβ dimers results in autoxidation, structural destabilization of heme within its binding pocket, and subsequent transfer of heme prosthetic groups to lipids, proteins and bacterial receptor complexes. In addition, acellular Hb and its byproducts can scavenge nitric oxide (NO) and increase oxidative stress, cause hypertension, vasoconstriction, kidney injury, and cardiovascular lesions.
In the circulatory intravascular compartment, acellular Hb and one of its byproducts, heme, are bound and cleared by the plasma proteins haptoglobin (Hp) and hemopexin (Hpx), respectively. The ability of Hp to control acellular Hb exposure is dependent on the primary binding site for Hb αβ dimers. Irreversible binding between the Hp β chain and Hb αβ dimers occurs at a 1:1 stoichiometric ratio. This interaction provides a critical antioxidant function by protecting key amino acids in Hb that are vulnerable to oxidation, preventing heme release and compartmentalizing Hb within a stable protein complex that does not extravasate out of the circulatory intravascular compartment into the parenchymal tissues. During hemolytic diseases, Hp acts as the primary defense, stabilizing acellular Hb. Upon Hp saturation the dimerized excess Hb releases heme, which is captured by Hpx. In addition to Hpx, serum albumin can act as a secondary heme scavenger. However, albumin's affinity for heme is lower than that of Hpx and the heme-albumin complex does not fully protect from heme-mediated toxicity.
Apohemoglobin (apoHb) is produced by removing heme from Hb. This apoprotein has a high affinity region in the unoccupied hydrophobic heme-binding pocket for a number of ligands, but the highest affinity of apoHb is for heme. Some of the prior examples have illustrated the binding of heme to apoHb and the complexation of apoHb to Hp in vitro, forming an apoHb-Hp complex. In pathophysiological states of heme excess, the use of apoHb or apoHb-Hp may offer an advantage by scavenging and clearing excess heme through the monocyte/macrophage CD163 surface receptor. However, no study has explored the potential in vivo heme scavenging properties of apoHb or apoHb-Hp. Moreover, it has been observed that the apoHb-Hp complex, in addition to heme-binding, can exchange Hp bound apoHb dimers for holoHb dimers, serving a dual role of Hb and heme scavenger in vivo (see
It was hypothesized that apoHb-Hp could facilitate heme transfer during states of high heme stress or holoHb exchange for apoHb dimers during acellular Hb exposure. Further, we hypothesized that these ligand interactions could attenuate the adverse microcirculatory consequences of heme and acellular Hb. In the present example, a sequential approach was tested for characterizing in vitro and in vivo apo-Hb-Hp and heme/Hb ligand interactions using a guinea pig model to assess time dependent heme transfer and Hb exchange. To test the effect of apoHb for holoHb exchange on vascular response, apoHb, Hp and apoHb-Hp complex pre-treatment followed by Hb administration as a 10% top-load was evaluated. Next, the effect of heme transfer to apoHb-Hp on the vascular response to heme-albumin was tested after apoHb-Hp as a 20% blood volume exchange. Both experimental designs were conducted in conscious Golden Syrian hamsters instrumented with the dorsal window chamber model to quantify systemic and microvascular hemodynamic responses to heme and acellular Hb.
Methods
Hb Preparation
Human Hb for this study was prepared via tangential flow filtration as described in the examples. Expired units of human red blood cells were generously donated. The concentration of Hb was determined spectrophotometrically. All Hb (also referred to as holoHb) used in this study was of human origin and derived via this method with over 97% oxyHb prior to use.
Apohemoglobin Preparation
The apoHb used in this study was prepared via tangential flow filtration based on the acidic-ethanol heme-extraction procedure as described in the examples. The absence of residual heme was verified by measuring the ratio of absorbance between the Soret peak (λmax=412−413 nm) and 280 nm to ensure less than 1% residual heme in the product. The heme-binding capacity of apoHb preparations was 80%.
Haptoglobin Preparation
Human Hp was purified from human Cohn fraction IV (FIV) purchased from Seraplex (Pasadena, Calif.). The final protein solution was composed of a mixture of Hp2-1 and Hp2-2 Hp polymers with an average MW of 400-500 kDa.
Heme-Albumin Preparation
A 4 mM heme-albumin stock solution was prepared by dissolving 65 mg Hemin (Sigma) in 100 mM NaOH at 37° C. and incubating for 1 h at 37° C. with 20% human serum albumin (HSA) and purchased from Grifols (Los Angeles, Calif.). The pH was carefully adjusted to 7.45 with 14 mM orthophosphoric acid/317 mM NaCl followed by sterile filtration.
Total Protein Assays
To estimate the total protein concentration of the apoHb solution, a Bradford assay was performed (Coomassie Plus Protein assay kit, Pierce Biotechnology, Rockford, Ill.). Additionally, spectrophotometric analysis of the 280 nm peak for each sample was used to estimate the total protein concentration using the millimolar extinction coefficients of apoHb found in the literature.
In Vitro Hb Exchange and Heme Binding by the ApoHB-Hp Complex
The apoHb-Hp complex was formed by reacting apoHb with Hp, and complex formation was confirmed by analysis of the mixture via size exclusion HPLC (HPLC-SEC). Heme binding to the apoHb-Hp complex was assessed by mixing the apoHb-Hp complex with heme-albumin prior to HPLC-SEC. Furthermore, apoHb exchange for Hb within the apoHb-Hp complex was determined by reacting the apoHb-Hp complex with Hb. HPLC-SEC was performed using an Acclaim SEC-1000 (4.6×300 mm) column (Thermo Fisher Scientific, Waltham, Mass.) attached to a Dionex UltiMate 3000 system (Thermo Fisher Scientific, Waltham, Mass.). The mobile phase consisted of 50 mM potassium phosphate, pH 7.4 at a flow rate of 0.35 mL/min controlled by Chromeleon 7 software. Wavelength absorbance used for monitoring the protein concentration was set to λ=280 nm, λ=405 nm to detect heme (bound to apoHb or albumin), and λ=413 nm to detect Hb, as it corresponds to the intermediate wavelength between residual heme in apoHb (λ=412 nm) and prevalent holoHb form used in the study, oxyHb (λ=415 nm).
In Vivo Hb Exchange and Heme Binding by the ApoHb-Hp Complex
In vivo heme transfer (heme-albumin) and Hb (acellular Hb) exchange with apoHb-Hp was studied in male Hartley guinea pigs (400-600 grams). Guinea pigs were infused with acellular Hb (0.75 μmol protein) (n=5) or heme-albumin (0.75 μmol protein) (n=5) followed by apoHb (0.75 μmol protein) bound to Hp. All materials were pre-filtered using a 0.22 pm syringe filter prior to injection. Plasma concentrations of all components were evaluated over a 6-hour period by UV-visible spectrophotometry and analytical HPLC-SEC. Plasma concentrations versus time are expressed as heme (μm/ml). On days of surgery, guinea pigs were dosed subcutaneously with ketoprofen (5 mg/kg) for pain management and then anesthetized via intraperitoneal injection with a cocktail of ketamine HCl (100 mg/kg) and xylazine HCl (5 mg/kg) (Phoenix Scientific Inc., St. Joseph, Mo. USA). Sterilized PESO tubing catheters were placed into the left external jugular vein, and left carotid artery and exteriorized at the back of the neck, all surgical sites were treated topically with bupivacaine HCl (2.5 mg/ml) (AuroMedics, Windsor, N.J., USA), and closed with 4-0 surgical silk internal sutures and external surgical staples. After 24-hours of post-surgical recovery, conscious guinea pigs were randomly allocated to dosing groups. Blood samples (150 μL) were collected through the carotid artery at baseline, after acellular Hb or heme-albumin albumin infusion, immediately after apoHb-Hp infusion (time 0) then at 0.25, 0.50, 0.75, 1.00, 1.25, 1.50, 3.00 and 6.00 hours.
Plasma Hemoglobin Analysis
Blood samples were centrifuged at 2,000 rpm for 10 min immediately after collection. Plasmas were diluted in 50 mM phosphate-buffered saline and analyzed on the same days of blood collection using a Carey 60 UV-visible spectrophotometer (Agilent Technologies, Santa Clara, Calif.). Oxy ferrous Hb [HbFe2+O2] and ferric Hb [HbFe3+] concentrations were determined based on the extinction coefficients for each species. Molar extinction coefficients used to calculate Hb concentrations in heme equivalents were: 15.2 mM−1 cm−1 at 576 nm for Hb(O2) and 4.4 mM−1 cm−1 at 631 nm for ferric Hb using 50 mM potassium phosphate buffer, pH 7.0 at ambient temperature, in both cases. Total heme was calculated by adding these values.
HPLC-SEC Anlysis of Hb-Hp Complexes in Plasma
Plasma samples (50 μL) were injected into a BioSep-SEC-S3000 (600 7.5 mm) SEC column (Phenomenex, Torrance, Calif.). The SEC column was attached to a Waters 2535 Quaternary Gradient Module pump and Waters 2998 Photodiode Array Detector controlled by a Waters 600 controller using Empower 2 software (Waters, Milford, Mass.), wavelength monitoring was the same as used for in vitro analysis (280, 405 and 413 nm).
Window Chamber Preparation in Golden Syrian Hamsters
Studies were performed in 55-65 g male Golden Syrian Hamsters (Charles River Laboratories, Boston, Mass.) fitted with a dorsal window chamber. The hamster window chamber model is widely used for microvascular studies without anesthesia. The complete surgical technique has been described previously. Two days after window implantation, arterial and venous catheters filled with heparinized saline solution (30 IU/mL) were implanted into the carotid and jugular vessels. Catheters were tunneled under the skin, exteriorized at the dorsal side of the neck, and securely attached to the window frame.
Inclusion Criteria
Hamsters were considered suitable for experiments if systemic parameters were as follows: heart rate (HR)>340 beats/min, mean arterial blood pressure (MAP)>80 mm Hg, systemic Hct>45%, and arterial 02 partial pressure (pAO2)>50 mm Hg. Additionally, hamsters with signs of low perfusion, inflammation, edema, or bleeding in their microvasculature were excluded from the study. Guinea pigs were included in the study if they were deemed healthy and met the weight range criteria of 400-600 g.
Experimental Setup
The unanesthetized animal was placed in a restraining tube with a longitudinal slit from which the window chamber protruded, then fixed to the microscopic stage for transillumination with the intravital microscope (BX51WI, Olympus, Japan). Animals were given 20 minutes to adjust to the tube environment and images were obtained using a CCD camera (4815, COHU, San Diego, Calif.). Measurements were carried out using a 40× (LUMPFL-WIR, numerical aperture 0.8, Olympus) water immersion objective.
Systematic Parameters
The MAP and HR were monitored continuously (MP150, Biopac System Inc., Santa Barbara, Calif.). Hct was measured from centrifuged arterial blood samples taken in heparinized capillary tubes. Hb content was determined spectrophotometrically (B-Hemoglobin, Hemocue, Stockholm, Sweden). Arterial blood was collected in heparinized glass capillaries (50 μL) and immediately analyzed for pO2, pCO2, and pH (ABL90; Radiometer America, Brea, Calif.). Arterial Hb saturation was measured using an IL482 CO-Oximeter (Instrumentation Laboratory, Lexington, Mass.).
Microhemodynamics
Arteriolar and venular blood flow velocities were measured using the photodiode cross-correlation method (Photo-Diode/Velocity, Vista Electronics, San Diego, Calif.). The measured centerline velocity (V) was corrected according to blood vessel size to obtain the mean RBC velocity. A video image-shearing method was used to measure blood vessel diameter (D). Blood flow (Q) was calculated from the measured values as Q=π/4D2 V.
Functional Capillary Density (FCD)
Functional capillaries, defined as capillary segments that have transit of at least one RBC in a 60 second period in 10 successive microscopic fields, were assessed in a region of 0.46 mm2. The FCD (cm−1) was calculated as the total length of RBC perfused capillaries divided by the viewing area (0.46 mm2).
Vascular Response to Acellular Hb
Hamsters were pre-treated with apoHb (30 mg/mL, 100 μL, n=6), Hp (15 mg/mL, 50 μL, n=6), apoHb-Hp complex (45 mg/mL, apoHb 15 mg/mL+Hp 30 mg/mL, 150 μL, n=6), or vehicle (saline equal volume as study groups, n=6). Hamsters were then subjected to a 10% hypervolemic infusion of purified acellular human Hb (50 mg/mL, 500 μL) and systemic and microvascular responses were measured after 30 minutes. All materials were pre-filtered using a 0.22 μm syringe filter prior to injection.
Vascular Response to Heme Albumin
Hamsters were either pre-treated with apoHb-Hp complex (45 mg/mL, apoHb 15 mg/mL+Hp 30 mg/mL-0.15 mL and 6.75 mg total dose, n=6), or vehicle (saline equal volume as study groups, n=6). Then hamsters were dosed with heme-albumin (2.0 mg/mL, 2.0-2.8 mg total dose) via a 20% blood volume exchange transfusion. All materials were pre-filtered using a 0.22 μm syringe filter prior to injection. Systemic hemodynamics and microcirculatory function were measured after 30 minutes.
Statistical Analysis
For in vivo Hb exchange and heme transfer studies performed in guinea pigs (n=5/group), all data are represented as mean values±SD. AUC values were derived using the linear trapezoidal rule and data were compared using a One-way ANOVA with Tukey's multiple comparisons test for parametric data in GraphPad Prism 8.3, GraphPad Software Inc., San Diego, Calif. For microcirculatory studies, data within each group were analyzed using a two-way ANOVA for repeated measurements. When appropriate, post hoc analyses were performed with the Dunnett's multiple comparisons test. Microhemodynamic data are presented as ratios relative to baseline values and absolute values are reported in the supplementary tables. A ratio of 1.0 signifies no change from baseline, while lower and higher ratios are indicative of changes proportionally lower and higher than baseline (i.e., 1.5 represents a 50% increase from the baseline level). The same blood vessels and capillary fields were monitored throughout the study, such that direct comparisons to their baseline levels could be performed, allowing for more reliable statistics on small sample populations. All statistics were calculated using GraphPad Prism 6 (GraphPad Software, Inc., San Diego, Calif.). Changes were considered significant if p values were less than 0.05.
Results
In Vitro HoloHb Exchange and Heme Binding by the ApoHb-Hp Complex
Formation of the apoHb-Hp complex in vitro can be achieved by reacting the two proteins (apoHb+Hp). The difference in molecular weight (MW) of Hp (˜400 kDa in this study) compared to apoHb (˜32 kDa) allowed for assessment of apoHb-Hp complex formation via HPLC-SEC. This is shown in the SEC chromatograms of
In Vivo HoloHb Exchange for ApoHb within the Hb-Hp Complex Guinea pigs were administered holoHb (denoted as Hb) followed by apoHb-Hp and time dependent blood sampling for analysis of Hp-bound and -unbound Hb plasma concentrations versus time are shown in
In Vivo Heme Transfer from Heme-Albumin to ApoHb within the ApoHb-Hp Complex
Guinea pigs were administered heme-albumin followed by apoHb-Hp. Time dependent blood sampling was conducted for analysis of heme transfer from heme-albumin to apoHb-Hp resulting in heme intercalated into apoHb αβ dimers, generating bound holoHb αβ dimers (denoted Hb-Hp) and their plasma concentrations over time as shown in
Microvascular Impact of Pretreatment with ApoHb, Hp and ApoHb-Hp Complex Followed by Acellular Hb Infusion
Systemic Parameters
The MAP and HR after pretreatment with vehicle, apoHb, Hp or apoHb-Hp, and subsequent challenge with acellular Hb dosing are presented in
Functional Capillary Density
The FCD after pretreatment with vehicle, apoHb, Hp or apoHb-Hp, and challenge with acellular Hb dosing are presented in
Microhemodynamics
The changes in arteriolar and venular hemodynamics relative to baseline are shown for all four experimental groups. The arterioles were segmented into various vessel diameters: small arterioles from 20 to 40 μm, mid-size arterioles from 40 to 60 μm and large arterioles from 60 to 100 μm. All venules were very consistent and grouped into a single venule group from 30 to 80 μm in vessel diameter.
Small Arterioles (20-40 μm): The diameter, velocity and flow, relative to baseline, for small arterioles, ranging from 20-40 μm in diameter are shown in
Mid-Sized Arterioles (40-60 μm): The diameter, velocity and flow, relative to baseline, for medium sized arterioles, ranging from 40-60 μm in diameter are shown in
Large Arterioles (60-100 μm): The diameter, velocity and flow, relative to baseline, for large arterioles, ranging from 60-100 μm in diameter are shown in
Venules (30-80 μm): The diameter, velocity and flow, relative to baseline, for venules, ranging from 30-80 μm in diameter are shown in
Taken together, these data suggest that apoHb-Hp may offer an improvement over Hp alone in microvascular function of small, mid-size and large arteriole. These data are potentially the first to demonstrate such an effect in the microcirculation and indicate the potential for a concomitant heme and Hb mediated pathobiology on the vasculature. To better understand this, the systemic and microcirculatory hemodynamic response to pre-treatment with our novel apoHb-Hp construct followed by heme-albumin exposure was studied.
In Vivo Pretreatment with ApoHb-Hp Followed by Heme-Albumin Exposure
This study was completed in twenty-four (N=24) instrumented animals. Eight animals were randomly assigned to each experimental group. The first experimental group was a control (untreated animals), and the second group was infused with apoHb-Hp. All groups were then administered a single dose of heme-albumin. The impact of apoHb-Hp on mean arterial blood pressure (MAP) and heart rate (HR) following heme-albumin challenge are presented in
Discussion
Acellular Hb has multiple pathophysiologic effects when released into the intravascular space during hemolysis. Hb tetramers (α2β2) dissociate into αβ dimers quickly in the circulation which can easily oxidize into methemoglobin (metHb) and release free heme. Multiple pathophysiological consequences are associated with heme and Hb. To prevent the adverse effects from Hb and its degradation products, Hp binds dimeric Hb (i.e. αβ dimers), thus preventing Hb from mediating oxidative reactions and from releasing free heme. When Hp is depleted (during extensive acute or low-level chronic hemolysis), heme released by Hb is transferred and bound by Hpx. In addition, albumin serves as a secondary heme binding protein, but does not fully prevent heme-mediated toxicity due to its lower affinity for heme than highly lipophilic LDL, HDL and VLDL. Hb binding to Hp is a biologically irreversible process, creating a stable and high-molecular-weight molecule that is cleared by macrophages and monocytes upon binding to the scavenger receptor, CD163. ApoHb acts as a heme scavenging protein, binding heme with a higher affinity than albumin. Moreover, the apoHb-heme protein can bind to Hp to be cleared through CD163+ macrophages and monocytes. To ensure apoHb-heme can be delivered to monocytes/macrophages apoHb was bound to Hp. This heme capture mechanism provides a specific pathway for heme clearance, in lieu of the described route of heme-Hpx uptake, which occurs through LRP-1 (low density lipoprotein receptor, CD91), a ubiquitous receptor that exists on the surface of numerous cell types and exhibits a multitude of functions.
This example demonstrates in vitro and in vivo holoHb exchange for apoHb when bound to Hp and highlights the unique property of heme intercalation into apoHb-Hp, following holoHb and heme-albumin exposure in guinea pigs. This combination of effects offers a potentially advantageous combination when concomitantly or independently addressing intravascular hemolysis and heme stress. Furthermore, microcirculatory studies in hamsters with apoHb, Hp, and the apoHb-Hp complex reduced adverse macrovascular and microvascular responses to acellular Hb exposure. More specifically, administration of Hp and apoHb-Hp complex prevented hypertension and vasoconstriction, as well as preserved microvascular diameter, blood flow, and functional capillary density across a range of arteriolar sizes. The apoHb-Hp complex appears to demonstrate a complementary mechanism to address Hb vasoactive response by attenuating heme and acellular Hb vascular interactions in the circulation. Unless bound to Hp, Hb dissociates into αβ dimers and extravasates through the blood vessel wall and reacts with or scavenges NO. Furthermore, both heme within Hb and heme bound to albumin participate in oxygen radical reactions that covalently modify proteins, lipids, carbohydrates, and nucleotides, leading to tissue damage. Administration of the apoHb-Hp complex may provide a unique therapeutic strategy to simultaneously mitigate heme stress and plasma Hb exposure across a range of disease states that involve heme-protein circulatory exposures. This approach provides a distinctive physiologic method to target the well-established mechanism of Hb-Hp clearance and intra-cellular Hb/heme detoxification by targeting the monocyte/macrophage CD163 surface receptor.
In the above microcirculatory function studies, infusion of acellular Hb induced arteriolar vasoconstriction and increased vascular resistance in small, mid and large arterioles. As Hb dissociates into αβ dimers it extravasates through the fenestrated capillaries and scavenges NO from the tissue. NO stimulates the production cGMP, decreasing the intracellular Ca2+ concentration, resulting in muscle relaxation; without the accumulation of cGMP the smooth muscles around the arterioles constrict. Hb mediated NO consumption leads to vasoconstriction, prevents perfusion of capillary beds, and reduces the number of functional capillaries, preventing metabolite washout. The vasoconstriction in the microcirculation results in an upstream response observed as an increase in MAP. The increase in MAP leads to a decrease in heart rate through the baroreceptor reflex. Finally, arteriole constriction hinders flow and velocity in all sized arterioles suggesting a reduction in tissue oxygen perfusion. The severity of these physiological responses can be observed after a simple exogenously administered acellular Hb intravascular dose. During active intravascular hemolysis, the body's natural levels of scavenger proteins (Hp and Hpx) cannot keep up with the demand to stabilize the Hb dimer or scavenge free heme and its associated iron.
In the above studies, a single injection of Hp, at a concentration of 1.5 g/dL, prior to the 0.1 mL Hb dose resulted a greater reduction in vasoconstriction compared to the control and the apoHb infused animals. This was an expected result, since Hp stabilizes the Hb αβ dimer and prevents the release of heme. Arterioles experienced less vasoconstriction, and flow as well as velocity were preserved to a greater degree. The downstream effect is that MAP and HR are maintained closer to baseline. Hp dosing also prevented the loss of FCD in response to heme and Hb, supporting the concept of intravascular compartmentalization which prevents extravasation of Hb through fenestrated capillaries, less NO scavenging, and preservation of microvascular pressures. For animals pretreated with 3 g/dL of apoHb, hemodynamic side-effects were observed in animals after Hb exposure. Most of the adverse hemodynamic responses were still present, including constriction of the arterioles, a decrease in velocity and flow, elevated systemic blood pressure, decreased heart rate and a reduction in functional capillaries. However, apoHb-Hp pre-dosing followed by Hb exposure lead to the most optimal response in terms of preserving systemic and microhemodynamic parameters close to the values found at baseline. ApoHb-Hp prevented the rise in MAP and decrease in HR. Further apoHb-Hp optimized blood vessel diameter, velocity and flow at baseline levels and the number of functional capillaries were maintained at basal levels. These results demonstrate a proof-of-concept that the apoHb-Hp complex prevents Hb-mediated circulatory dysfunction. In addition, apoHb-Hp was effective at maintaining basal circulatory function when dosed prior to heme-albumin exposure.
Conclusion
The present data suggests that the apoHb-Hp complex can effectively exchange apoHb for holoHb in vitro and in vivo. Further, apoHb situated in the apoHb-Hp complex is an effective heme binding protein complex based on in vitro and in vivo heme-albumin transfer studies and in vivo microcirculatory experiments. These unique properties of the apoHb-Hp complex prevent adverse systemic and microvascular responses to Hb and heme-albumin exposure and introduces a novel therapeutic approach to facilitate simultaneous removal of extracellular Hb and heme.
Polymerized hemoglobin (PolyHb) is a promising hemoglobin (Hb)-based oxygen (O2) carrier (HBOC) currently undergoing development as a red blood cell (RBC) substitute. Unfortunately, commercially developed products are comprised of low molecular weight (MW) PolyHb molecules, which extravasate, scavenge nitric oxide (NO), and resulted in vasoconstriction and hypertension. The naturally occurring Hb scavenging species, haptoglobin (Hp), combined with the purified heme scavenging species, apohemoglobin (apoHb), is a potential candidate to alleviate the pressor effect of PolyHb. In this example, the protective activity of administration of the apoHb-Hp complex was evaluated to mitigate the vasoactive response induced by the transfusion of low MW PolyHb. Hp binding to PolyHb was characterized in vitro. The effectiveness of apoHb-Hp administration on reducing the vasoconstriction and pressor effects of PolyHb was assessed by measuring systemic and microcirculatory hemodynamics. Transfusion of a low MW PolyHb to vehicle control pretreated animals increased mean arterial pressure (MAP), and decreased arteriolar dimeter, and functional capillary density (FCD). Transfusion of a low MW PolyHb to apoHb-Hp pretreated animals prevented changes in MAP, heart rate, arteriole diameter and blood flow, and FCD relative to before transfusion. These results indicate that the increased size of PolyHb after binding to the apoHb-Hp complex may help compartmentalize PolyHb in the vascular space and thus reduce extravasation, NO scavenging, and toxicity responsible for vasoconstriction and systemic hypertension.
Introduction
During the recent outbreak of severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), closure of blood donation centers and large exclusions in blood donor pools has resulted in severe blood shortages. A 2017 computer simulation of an influenza outbreak in the United States predicted that over 541,000 units of blood might be lost at the end of a 48-week influenza pandemic with a 65% reduction in blood donation rates. However, current red blood cell (RBC) donation trends, show that the donation rate decay due to SARS-CoV-2 is more severe than the previous influenza models. This deficit in the blood supply emphasizes the need for a RBC substitute when RBC units are not available. Unfortunately, there are currently no RBC substitutes approved for clinical application.
The presence of low molecular weight (MW) hemoglobin (Hb) species in previous generations of Hb-based oxygen (02) carriers (HBOCs) restricted their application as RBC substitutes. Among the myriad of strategies that can produce HBOCs, using glutaraldehyde crosslinking to form polymerized Hb (PolyHb) remains popular due to its low cost and excellent scalability. However, the commercially developed PolyHbs, HBOC-201 (Biopure Corp, Cambridge, Mass., USA) and PolyHeme (Northfield Laboratories Inc., Northfield, Ill., USA), primarily contained fractions of material at or below 250 kDa. Low MW PolyHb and unpolymerized Hb extravasate into the interstitial space where they scavenge nitric oxide (NO), resulting in vasoconstriction and systemic hypertension. Moreover, free heme release from HBCOs can also lead to systemic toxicity. Previously, it was determined that increasing the MW of PolyHb by increasing the molar ratio of glutaraldehyde to Hb attenuated hypertension and renal injury. However, low MW polymers comprise up to 40% of these PolyHb solutions. Unfortunately, removal of these low MW polymers would significantly reduce the yield of PolyHb. Alternatively, recent strategies to mitigate hypertension have used adenosine and nitroglycerine to attenuate the hypertensive response of HBOC-201. While effective at controlling systemic hypertension, administration of adenosine and nitroglycerine must be carefully controlled to prevent hypotension.
Instead, it is proposed that employing naturally occurring mechanisms of Hb detoxification is a potential strategy to mitigate systematic hypertension resulting from the presence of low MW PolyHb in the circulation. The naturally occurring Hb scavenging protein, haptoglobin (Hp), has a pivotal role in detoxifying stroma free Hb in the blood. A previous study demonstrated that Hp administration normalizes vascular NO signaling after hemolysis. Recently, it was determined that Hp preferentially binds to low MW PolyHb (MW<256 kDa). By binding to Hp, the MW of low MW PolyhHb is effectively increased. This increase in molecular size may reduce tissue extravasation and subsequent NO scavenging by PolyHb. An illustration of this potential detoxification mechanism is shown in
The second component of stroma-free Hb detoxification is hemopexin (Hpx). Hpx scavenges heme that is released from stroma free Hb after Hb auto-oxidation. By isolating heme within the Hpx complex, the heme is unable to catalyze oxidative reactions with blood and tissue components, thus preventing lipid, protein, and nucleic acid oxidation. A potential low-cost alternative to Hpx is heme-free apohemoglobin (apoHb). This molecule is able to scavenge free heme due to the highly hydrophobic nature of its vacant heme-binding pocket. Combining apoHb with Hp results in a protein complex (apoHb-Hp) that may be able to scavenge both Hb and heme in the plasma.
Because of its ability to compartmentalize stroma-free Hb and scavenge free heme, it was hypothesized that administration of the apoHb-Hp complex could reduce hypertension that results from the transfusion of low MW PolyHb. Hp binding to low MW PolyHb was confirmed in vitro with stop-flow fluorescence spectrometry and size exclusion chromatography. Mean arterial blood pressure (MAP) was used as an indicator of reduced hypertension. Additionally, intravital microscopy was used to examine how administration of apoHb-Hp influences functional capillary density (FCD), vascular tone, and blood flow after PolyHb administration.
Methods
PolyHb Synthesis and Biophysical Properties
The Hb used in this exampler was purified from human RBCs via tangential flow filtration (TFF), as described previously. Hb was polymerized with glutaraldehyde under complete deoxygenation in the tense (T) quaternary state at a 25:1 molar ratio of glutaraldehyde to Hb. After polymerization, PolyHb was clarified, purified, and buffer exchanged into a modified Ringer's lactate solution using TFF. Modified polyester sulfone hollow fiber filters with a molecular weight cutoff (MWCO) of 100 kDa were used to remove unpolymerized Hb from solution. The PolyHb solution was concentrated to 12 g/dL. The cyanomethemoglobin method was used to measure the Hb concentration and the metHb level of Hb/PolyHb solutions. The size distribution of PolyHb, by particle volume, was measured using dynamic light scattering (DLS) (Brookhaven Instrument Inc. BS-200M, Holtsville, N.Y.). The rheology of PolyHb solutions was measured using a DV3T-CP cone and plate rheometer (Brookfield AMETEK, Middleboro, Mass.) with cone spindle CPA-40Z. The O2-Hb/PolyHb equilibrium binding curves were measured using a Hemox Analyzer (TCS Scientific Corp., New Hope, Pa.). The Hb/PolyHb kinetics of O2 offloading (koff,O
ApoHb Purification and Biophysical Properties
The apoHb used in this study was produced using acidified-ethanol coupled TFF, as described previously. The heme-binding site activity of the resulting apoHb was quantified with a dicyanohemin assay. The total amount of protein in solution was estimated based on the molar extinction coefficient of apoHb.
Hp Purification and Biophysical Properties
The Hp used for this study was purified from human Cohn Fraction IV derived from pooled human plasma. The resulting Hp contained both Hp2-1 and Hp2-2 phenotypes. The total amount of protein in solution was estimated with a Bradford assay. The Hb binding capacity of Hp was assessed by monitoring Hb binding to Hp at 413 nm with HPLC-SEC and was used to quantify the concentration of Hp. The reaction of Hp with either unmodified human Hb or PolyHb synthesized in this study was monitored using an SX-20 stopped stopped-flow spectrophotometer using previously described methods (Applied Photophysics, Leatherhead, UK). Excess Hp (2:1, Hp:PolyhHb on a Hb tetramer binding basis) was allowed to react with PolyHb to form the resulting Hp-PolyHb complex. The resulting changes in size distribution after Hp binding was estimated with HPLC-SEC using previously described methods. All measurements were taken from the same sample used in the animal study.
Animal Model
In vivo evaluation of apoHb-Hp detoxification of PolyHb was performed on 55-65 g male Golden Syrian Hamsters (Charles River Laboratories, Boston, Mass.) fitted with a dorsal skinfold window chamber as previously described. Animals were considered suitable for experiments if systemic parameters were as follows: heart rate (HR)>340 beats/min, mean arterial blood pressure (MAP)>80 mm Hg, systemic Hct>45%, and arterial O2 partial pressure (pAO2)>50 mm Hg. Additionally, animals with signs of low perfusion, inflammation, edema, or bleeding in their microvasculature were excluded from the study. Prior to treatment, baseline systemic parameters and microvascular hemodynamics were assessed. Animals were first treated with either (1) 0.1 mL 0.9 wt % saline, (2) 0.1 mL apoHb (24.5 mg/L), or (3) 0.1 mL apoHb (24.5 mg/L) with 0.05 mL Hp (46.5 mg/mL). 20 minutes following the initial treatment, the systemic parameters and microvascular hemodynamics were assessed. Afterward, the animals underwent a 20 percent isovolemic exchange transfusion with a 10 g/dL PolyHb solution. After another 20 minutes, the systemic parameters and microvascular hemodynamics were measured. Mean arterial pressure (MAP) and heart rate (HR) were monitored continuously (MP150, Biopac System Inc., Santa Barbara, Calif.). The sample size of 5 animals per group was calculated based on an expected 10% difference in mean arterial pressure between groups, an α=0.05, and 1-β=0.1, and equal enrollment for all groups. Additionally, microhemodynamic measurements contain data from multiple vessels within the field (5-7 arterioles and venules selected at baseline based on visual clarity), improving the power of these measurements. Each experiment is a repeated measures study, so all experimental timepoints are replicated between all animals and groups. Furthermore, each experimental group contains animals from different litters, improving the replication of these studies. Animals were randomly assigned to their respective group before baseline measurements were taken. Each group contains animals from multiple litters of hamsters to improve randomization. Investigators were not blinded to group allocation during data collection or analysis. Blinding is not possible with these solutions as they have distinct colors. Measurements taken are highly quantitative, so blinding should have little impact.
Intravital Microscopy
The unanesthetized animal was placed in a restraining tube with a longitudinal slit from which the window chamber protruded, then fixed to the microscopic stage for transillumination with the intravital microscope (BX51WI, Olympus, New Hyde Park, N.Y.). Animals were given 20 minutes to adjust to the tube environment and images were obtained using a CCD camera (4815, COHU, San Diego, Calif.). Measurements were carried out using a 40× (LUMPFL-WIR, numerical aperture 0.8, Olympus) water immersion objective.
Microvascular Hemodynamics
Arteriolar and venular blood flow velocities were measured using the photodiode cross-correlation method (Photo-Diode/Velocity, Vista Electronics, San Diego, Calif.). The measured centerline velocity (V) was corrected according to blood vessel size to obtain the mean RBC velocity. A video image-shearing method was used to measure blood vessel diameter (D). Blood flow (Q) was calculated from the measured values as
Functional Capillary Density (FCD)
Functional capillaries, defined as capillary segments that have RBC transit of at least one RBC in a 60 second period in 10 successive microscopic fields, were assessed in a region of 0.46 mm2. FCD (mm−1) is calculated as the total length of RBC perfused capillaries divided by the area (0.46 mm2).
Statistical Analysis
Results are presented as mean±standard deviation. One-way ANOVA was used to analyze data within the same group. All box plots are presented with the median on the center line, the box limits are set to the upper (75%) and lower (25%) quartile. All outliers are shown each plot. Post-hoc analysis was completed with the Dunn multiple comparison test. Data between groups were analyzed with a two-way ANOVA with Bonferroni tests. When possible, in vivo data was compared against baseline in the same animal or same vessel as a ratio relative to the baseline. All statistical calculations and data analyses were performed with R (v. 3.6.2). For all tests, P<0.05 was considered statistically significant. All data is available upon reasonable request.
Results
PolyHb Characterization
A summary of the biophysical properties of the PolyHb used in this study is shown in
Characterization of Hp Binding to PolyHb
The effect of Hp binding on the size distribution of PolyHb and Hb is shown in
Systemic Hemodynamics
MAP and HR measured throughout the experimental study are shown in
Microhemodynamics
Changes in arteriole and venule diameter as measured with intravital microscopy are shown in
Changes in the blood velocity and volumetric flow rate measured via intravital microscopy are shown in
In the apoHb-Hp treatment group, there were no significant changes in arteriole and venule fluid velocities and volumetric flow rates throughout the study. The venule fluid velocity in the apoHb-Hp treatment group was significantly lower than the saline group and significantly higher than the apoHb treatment group. In the apoHb-Hp group, the arteriole and venule volumetric flow rates after PolyHb administration were significantly higher compared to the saline and apoHb treatment groups.
Changes in FCD throughout the intravital microscopy study are displayed in
Discussion
The principal finding of this example is that administration of a Hb and heme scavenging material (apoHb-Hp) maintained hemodynamics after a 20% isovolemic exchange transfusion with a low MW PolyHb. In animals administered with apoHb-Hp, there were negligible changes in MAP, HR, and microhemodynamics compared to baseline conditions. When compared to the systemic and microcirculatory changes observed in the apoHb and saline groups, the relatively small changes in the apoHb-Hp group indicate that Hp-based species may serve as appropriate materials to counteract the vasoactive effects of low MW HBOCs that are capable of binding to Hp.
The biophysical properties of the Hp produced in this study was comparable with values measured in the literature. The rate constant for Hp binding to Hb was similar to values reported in the literature for Hb (0.129 μM−1s−1). The rate constant for Hp binding to 25:1 PolyHb was much higher than the values reported in the literature for a PolyHb (0.003 μM−1s−1) and Oxyglobin (0.011 μM−1s−1). However, it was comparable to previously produced PolyHb. Even though the rate of Hp binding to PolyHb was significantly lower than the rate of Hp binding to Hb, systemic and microcirculatory hemodynamics were maintained throughout the studies. This indicates that Hb does not significantly inhibit Hp binding to PolyHb. By observing the change in fluorescence intensity, we were able to calculate the percentage of PolyHb that is capable of binding to Hp. For Hb, we calculated complete binding site saturation when excess Hb was in solution. In contrast, only a fraction of the binding sites was saturated after excess PolyHb was added to the Hp solution. Despite only observing fractional binding site saturation, dramatic increases in the MW and diameter of PolyHb after the Hp was added were still observed. This indicates that Hp is likely capable of binding higher order (MW>64 kDa) PolyHb molecules. Thus, Hp likely has a role in compartmentalizing 0, 1st, and 2nd order PolyHbs in the circulation.
The O2 affinity of the low MW PolyHb used in this example (31.1±0.5 mm Hg) was similar to the O2 affinity of Hb in human RBCs (P50=29.3). While the P50 was similar to Hb in RBCs, the lack of cooperativity (n=0.98±0.2) likely results in changes to O2 offloading in the arterioles. The O2 affinity was also comparable to previously produced PolyHbs (P50=30.7±1.2 mm Hg) and PolyHeme (P50˜29 mm Hg). Despite the fact that the average MW and diameter of the PolyHb produced for this example (AVG MW: 480 kDa), is significantly greater than previously produced commercial products [PolyHeme (64-400 kDa, AVG: 150 kDa) and HBOC-201 (69-500 kDa, AVG: 250 kDa)3], the PolyHbs produced in this example are comprised of approximately 50% low MW species (MW<500 kDa), which are known to be vasoactive than the higher MW species.
The relative changes in MAP and vessel tone compared to baseline in the saline treatment group were, on average, comparable to the changes observed after a top-load dose of HBOC-201. This is expected, given that the low MW PolyHb molecules used in this example had similar size distributions and O2 affinity compared to HBOC-201. The relatively similar properties of the PolyHb produced make it a promising surrogate for previous commercial products.
Administration of apoHb alone had relatively little effect on maintaining hemodynamics after transfusion of PolyHb. In many cases, apoHb administration resulted in similar changes relative to baseline compared to the saline treatment group. This is expected given that the rate of heme release from PolyHbs is relatively low compared to stroma free Hb. More importantly, since apoHb exists as an αβ dimer (32 kDa), it was expected to be rapidly cleared through the kidneys within 5 minutes to an hour after administration. Moreover, heme binding to apoHb yields Hb which can cause similar effects as low MW PolyHb. However, the circulatory half-life would be increased if the apoHb were to bind plasma Hp. In addition, heme-binding to apoHb bound to Hp would neutralize the effects of free heme. Interestingly, the HR significantly decreased after PolyHb transfusion in the saline treatment group. This effect did not occur in the groups administered with the apoHb or apoHb-Hp solutions.
In contrast to both apoHb and saline, the apoHb-Hp complex was successful at maintaining baseline hemodynamics after transfusion of PolyHb. This indicates that Hp likely can mitigate the pressor effect of PolyHb by compartmentalizing low MW PolyHb within the vascular lumen. This mechanism is likely similar to the previously reported mechanism of cell-free Hb localization which preserved NO signaling. By increasing the MW of PolyHb via exchange of apoHb in the apoHb-Hp complexes with PolyHb, and thus eliminating the presence of small PolyHb species, translocation of PolyHb may be effectively stopped.
Previous studies have attempted to counteract the pressor effect of HBOCs via coadministration of adenosine or nitroglycerin. However, these materials require careful control of the dose to avoid hypotension. In contrast, independent administration of the apoHb-Hp solution did not significantly decrease MAP or alter HR compared baseline conditions. Unlike methods that target NO or endothelin, apoHb-Hp scavenging directly targets low MW stroma free Hb species. By targeting the Hb species directly, there is no need to balance a pressor response.
In conclusion, the results of this example indicate that the apoHb-Hp complex is a promising biomaterial that may make HBOCs safer for clinical application via mitigation of the pressor effect.
Photodynamic therapy (PDT) has been shown to effectively treat cancer by producing cytotoxic reactive oxygen species (ROS) via excitation of photosensitizers (PS). However, most PS lack tumor cell specificity, possess poor aqueous solubility, and cause systemic photosensitivity. Removing heme from hemoglobin (Hb) yields an apoprotein called apohemoglobin (apoHb) with a vacant heme-binding pocket that can efficiently bind to hydrophobic molecules such as PS. In this example, the PS aluminum phthalocyanine (Al-PC) was bound to the apoHb-haptoglobin (apoHb-Hp) protein complex, forming an apoHb-Al—PC-Hp (APH) complex. The reaction of Al-PC with apoHb prevented Al-PC aggregation in aqueous solution, retaining the characteristic spectral properties of Al-PC. The stability of apoHb-Al-PC was enhanced via binding with Hp to form the APH complex which allowed for repeated Al-PC additions to maximize Al-PC encapsulation. The final APH product had 65% of the active heme-binding sites of apoHb bound to Al-PC and hydrodynamic diameter of 18 nm, that could potentially reduce extravasation of the molecule through the blood vessel wall and prevent kidney accumulation of Al-PC. Furthermore, more than 80% of APH's absorbance spectra was retained when incubated for over a day in plasma at 37° C. Heme displacement assays confirmed Al-PC was bound within the heme-binding pocket of apoHb and binding specificity was demonstrated by ineffective Al-PC binding to human serum albumin, Hp, or Hb. In vitro studies confirmed enhanced singlet oxygen generation of APH over Al-PC in aqueous solution and demonstrated effective PDT on human and murine cancer cells. Taken together, this example provides a method to produce APH for enhanced PDT via improved PS solubility and potential targeted therapy via uptake by CD163+ macrophages and monocytes in the tumor (i.e. tumor associated macrophages). Moreover, this scalable method for site-specific encapsulation of Al-PC into apoHb and apoHb-Hp may be used for other hydrophobic therapeutic agents.
Introduction
Photodynamic therapy (PDT) is a treatment modality that generates reactive oxygen species (ROS) to induce localized cell death by either apoptosis or necrosis. PDT generates ROS by exciting photosensitizer (PS) molecules, in which the irradiated PS stimulates an electron to an unstable excited state. The energy from this electron can be transferred to an organic molecule (type I) or directly to molecular oxygen (type II). The former creates radicals which react with oxygen to form ROS, while the later directly forms singlet oxygen (1O2), a specific form of ROS. Even though both mechanisms lead to cell death, the type of damage varies. It has been shown that type I leads to cell death via necrosis, while type II leads to apoptosis. In addition to inducing tumor cell death, PDT has been shown to disrupt the tumor vasculature, and induces an anti-tumor immune response (potentially capable of preventing metastasis). PDT is a minimally invasive technique that has greatly expanded its potential biomedical applications with the development of fiber-based light delivery systems. Moreover, not only has PDT already shown great potential against cancer, but the combination of PDT with other cancer treatment modalities such as surgery or radiotherapy has shown synergistic effects with no cross-resistance. Further, PDT can be used in non-cancerous diseases such as age-related macular degeneration and as an antimicrobial agent.
For successful PDT treatment, potent PS molecules are required. First generation PS's exhibited low ROS production and prolonged photosensitivity, limiting their clinical use. Thus, researchers began developing new PS's for PDT. Phthalocyanines (PCs) are a low cost, second generation PS with high rate of ROS production and improved tissue penetration. PC's are stable and easily synthesized with low toxicity. Additionally, metalized-PCs can induce ROS generation via an alternative pathway by catalyzing the Fenton and Haber-Weiss reactions; as well as, significantly reducing levels of the antioxidant glutathione. One promising PS of this class is aluminum-PC (Al-PC) which has already shown promise as a PDT agent in previous studies and is clinically approved for use in Russia for PDT of stomach, lung, skin, lip, esophagus, oral cavity, tongue, and breast cancers as well as age-related macular degeneration. However, the Russian Al-PC comes in a mixture of sulphonated Al-PC which, although it improves aqueous solubility, has been shown to decrease photodynamic activity.
Even though new potent PS molecules have been developed, treatment has mainly relied on the unspecific tendency of particles to accumulate in tumor cells (attributed to the leaky and tortuous blood vessels of tumors). Furthermore, current applications of PDT are limited by the PS's low aqueous solubility and tendency to aggregate. Thus, even though there has been progress in the field, there is a need for more efficient delivery methods. Some examples of improved delivery include binding to albumin or low-density lipoprotein conjugates, targeting moieties (steroids, peptides, antibodies), liposomal encapsulation and nanocarriers. However, these mechanisms still lack tumor targeting, can lose PS via transfer to serum particles, or have complicated and expensive development. These issues lead to systemic and prolonged photosensitivity in patients, restricting the application of PDT. Moreover, new proposed delivery systems have been able to transport large quantities of PS molecules, but the PS molecules aggregated within the transport vehicle as indicated by the low absorbance of the species.
A promising candidate for transport and delivery of PS molecules is apohemoglobin (apoHb). Prosthetic heme groups are tightly bound inside the hydrophobic heme-binding pocket of hemoglobin (Hb) that can be removed to yield apoHb. The vacant heme-binding pocket of apoHb facilitates binding of hydrophobic molecules to the apoprotein, thus creating an aqueous drug delivery vehicle for hydrophobic molecules. For example, in Hb, apoHb binds to heme, enhancing heme's aqueous solubility and preventing its aggregation so that oxygen can be bound to the heme groups and transported via the circulatory system. Moreover, the use of apoHb for drug delivery has the benefit that the globin structure of apoHb resembles that of Hb and should exhibit little to no immune response. In addition to potentially serving as a drug carrier, the apoprotein clears via the same pathway as cell-free Hb in which CD163+ macrophages and monocytes uptake the protein after haptoglobin (Hp) binding. Due to this specific uptake mechanism, Hb and apoHb-based drug delivery systems have already been used to target CD163+ macrophages and monocytes. Based on these characteristics, apoHb is a promising yet simple targeted drug delivery vehicle for small hydrophobic molecules such as PS molecules.
In this example, the solubility and delivery challenges associated with PS molecules were addressed by using apoHb bound to Hp as a carrier of Al-PC. ApoHb was complexed with the plasma protein Hp to improve the apoprotein's stability in vivo and increase binding of Al-PC to the protein. Hp is the primary scavenger of cell-free Hb and delivering it to CD163+ macrophages and monocytes for recycling into CO, iron and biliverdin. Binding of apoHb-Al-PC to Hp also leads to the formation of a large apoHb-Al—PC-Hp (APH) complex which can prevent extravascular extravasation of apoHb-Al-PC. Furthermore, ex vivo Hp binding to apoHb-Al-PC does not require depletion of endogenous plasma Hp for targeting to CD163+ macrophages and monocytes. Targeting to the CD163 receptor is a promising approach for cancer therapy, since CD163+ macrophages (M2 phenotype) are classified as tumor promoting and are found in high concentration in various tumors.
Targeting tumor associated macrophages (TAMs) is highly relevant due to their role in tumor promotion, metastasis, and immunotherapy. Moreover, TAMs are one of the most abundant cell types in tumors, accounting for approximately 50% of the tumor mass and consisting primarily of the M2 phenotype. Further, TAM targeting has been shown to serve as a drug depot for slow drug release into the tumor microenvironment. Finally, through selective targeting of M2 TAM, the APH complex could help identify the primary and metastatic tumors, potentially serving as theranostic molecule. Thus, the apoHb-Hp complex could serve as a targeted delivery system of therapeutic and imaging agents such as Al-PC for cancer theranostics.
A general diagram of the process used to synthesize APH is shown in
Materials and Methods
Materials. Sodium phosphate dibasic, sodium phosphate monobasic, aluminum phthalocyanine chloride, and hemin chloride were purchased from Sigma Aldrich (St. Louis, Mo.). Potassium cyanide, hydrochloric acid, sodium chloride, potassium chloride, 0.22 μm nylon syringe filters, and dialysis tubing (pore size: 6-8 kDa) were purchased from Fisher Scientific (Pittsburgh, Pa.). 0.2 μm Millex-GP PES syringe filters were purchased from Merck Millipore (Billerica, Mass.). Ethanol was purchased from Decon Labs (King of Prussia, Pa.). A KrosFlo® Research II tangential flow filtration (TFF) system and hollow fiber (HF) filter modules were obtained from Spectrum Laboratories (Rancho Dominguez, Calif.). Expired units of human red blood cells (RBCs) and thawed human plasma were generously donated by the Transfusion Service in the Wexner Medical Center at The Ohio State University (Columbus, Ohio). Human fraction IV paste was purchased from Seraplex, Inc (Pasadena, Calif.).
Fluorescence and Absorbance Spectroscopy. Ultraviolet-visible spectrometry was performed in quartz cuvettes using a HP 8452A diode array spectrophotometer (Hewlett Packard, CA) and fluorescence spectrometry was measured using a PTI fluorometer (Horiba Scientific, NJ).
Hb Preparation. Human Hb was purified via tangential flow filtration (TFF) as described previously. Hb concentration was determined spectrophotometrically via the Winterbourn equations.
ApoHb Production. Apohemoglobin was produced via TFF as described previously.
Hp Purification. Hp was purified from human Cohn fraction IV. The final protein solution was composed of a mixture of Hp2-1 and Hp2-2 Hp polymers with an average MW of 400-500 kDa.
Hb/ApoHb Binding Capacity of Hp. The binding capacity of Hp to Hb and apoHb was determined using size exclusion high performance liquid chromatography (HPLC-SEC). The change in the area under the curve of free Hb or apoHb chromatograms when Hp was added to the sample was used to quantify the amount of Hb or apoHb bound to Hp.
Total Protein Assay. The total protein concentration of apoHb in solution was measured using the molar extinction coefficient of apoHb at 280 nm (12.7 mM−1cm−1).
ApoHb Activity Assay. The activity of the vacant hydrophobic heme-binding pocket of apoHb was determined via the dicyanohemin (DCNh) incorporation assay as previously developed. The extinction coefficients of DCNh and rHbCN used were 85 mM−1 cm−1 and 114 mM−1 cm−1 at 420 nm, respectively.
Preparation of Al-PC solutions. Stock Al-PC samples were freshly made by dissolving 1 mg of Al-PC in 2 mL of 100% EtOH. The stock solution was then diluted in EtOH prior to addition into apoHb or apoHb-Hp samples in phosphate buffered saline (PBS, 0.1 M, pH 7.4). All samples with Al-PC were kept at 4° C. and wrapped in aluminum foil. The concentration of Al-PC solutions was determined via the extinction coefficient of 294 mM−1 cm−1 at 672 nm in EtOH.
Optimization of EtOH Addition to ApoHb. The stock Al-PC solution was diluted 100× into 100% EtOH. The diluted stock Al-PC was then added in increasing volumes to a constant volume of apoHb in PBS at a concentration of approximately 23 μM. Volume ratios of diluted stock Al-PC in EtOH to apoHb in PBS ranged from 1:40 to 1:4.44.
Stability of ApoHb-Al-PC. The stability of apoHb-Al-PC was assessed by adding 1 mL of Al-PC solution at ˜8 μM to 10 mL of apoHb in PBS at a concentration of ˜23 μM. The sample was then dialyzed against PBS using 6-8 kDa cellulose dialysis membranes to remove any residual EtOH. Dialysis was performed over three days with daily exchange of the PBS buffer. The UV-visible absorbance spectra of samples were measured before and after dialysis.
Optimization of Al-PC Loading to ApoHb Varying dilutions of the stock Al-PC solution were prepared in EtOH (2.78, 7.19, 9.86, 13.2, 16.6, 18.6, 20.1 μM). 200 μL of these solutions were individually added to 2 mL of apoHb in PBS at a concentration of approximately −23 μM. The absorbance spectra of the resulting mixtures were then measured via UV-visible spectrophotometry.
Production of APH Complex. APH was produced via a repeated addition protocol. Starting with 200 mL of apoHb at a concentration of ˜23 μM, 20 mL of Al-PC in EtOH at ˜18 μM was added dropwise into the apoHb solution under constant stirring. Hp was then added at equimolar concentration to bind to apoHb-Al-PC. The mixture was left to react for 1 hour at 4° C. EtOH was removed from the solution via TFF by diafiltration with PBS over a 50 kDa modified polyethersulfone (mPES) hydrophilic TFF filter with inlet pressure maintained at about 6.5 psig. The sample container and process tubing were wrapped with aluminum foil to prevent Al-PC exposure to light. After 6 diafiltration volumes, 2 mL of the product was sampled, and its absorbance spectra was measured. Then, 200 μL of the 18 μM Al-PC stock solution was added to the 2 ml sample from the system to test for further Al-PC binding to apoHb-Hp. If the absorbance peak at 680 nm increased, an additional 20 mL of the 18 μM Al-PC was added to the system via an injection port. This process was repeated until the absorbance at 680 nm did not increase with further Al-PC addition. The sample was then concentrated to approximately 2 AU/cm at 680 nm (˜1.3 mg/mL of APH) and stored at −80° C. with containers wrapped in aluminum foil. A general setup of the process along with a flowchart for the repeated Al-PC additions is shown in
APH Stability at 37° C. The stability of the apoHb-Al—PC-Hp complex (APH) was assessed in PBS and thawed human plasma at 37° C. APH was diluted four times into sealed cuvettes that were incubated at 37° C. and covered in aluminum foil for a total of 48 h. The absorbance spectra of samples were periodically measured to determine the loss of Al-PC from APH over time. The fraction retained was determined by the ratio of the absorbance at 680 nm compared to the initial absorbance at 680 nm. The effects of incubation time, plasma exposure and the interaction of incubation time and plasma exposure were determined via a mixed-model approach using JMP Pro 13.
Specificity of Al-PC Binding to ApoHb. Binding of Al-PC to human serum albumin (HSA), Hb, reconstituted Hb (rHb), and Hp was tested to assess if binding to Al-PC was specific to the vacant heme-binding pocket of apoHb. Binding studies were performed via the same protocol described in Optimization of EtOH Addition to ApoHb. The concentrations of Hp, HSA, Hb and rHb in solution were 1.0 mg/mL, 0.35 mg/ml (and 3.5 mg/ml), 0.35 mg/mL, and 0.33 mg/mL respectively.
Heme-Albumin Mediated Displacement of Al-PC from APH. A fixed APH concentration (˜0.4 mg/mL) was mixed with increasing concentrations (0, 1.5 and 7.5 μM) of heme-albumin. Samples were kept isolated from light and incubated at both 4° C. and 37° C. Absorbance spectra of the mixtures were periodically measured to determine the displacement of Al-PC from apoHb in the APH complex via a decrease in the 680 nm peak, and transfer of heme from heme-albumin to apoHb in the APH complex via an increase in the 405 nm peak. Pseudo-first order rate constants were determined, which were then adjusted for the concentration of heme-albumin to determine the second order rate constant of Al-PC displacement by heme.
Dynamic Light Scattering. The hydrodynamic diameter of samples was determined using a BI-200SM goniometer (Brookhaven Instruments, Holtsville, N.Y.) at an angle of 90° and wavelength of 637 nm. Samples ware diluted to ˜1 mg/mL concentration in PBS (0.1 M, pH 7.4). The hydrodynamic diameter was calculated from experimental data by using the non-linear least squares (NNLS) algorithm in the instrument software.
HPLC Size Exclusion Chromatography. Samples were separated on an analytical Acclaim SEC-1000 (4.6×300 mm) column (Thermo Fisher Scientific, Waltham, Mass.) attached to a Dionex UltiMate 3000 system (Thermo Fisher Scientific, Waltham, Mass.). The mobile phase consisted of 50 mM potassium phosphate, pH 7.4. The flow rate and UV-visible spectral detection was controlled on Chromeleon 7 software with detection set to λ=280 nm and λ=680 nm to detect protein and Al-PC elution respectively at a flow rate 0.35 mL/min.
Singlet Oxygen Detection. Singlet oxygen was detected using the probe 1,3-diphenylisobenzofuran (DPBF), which reacts irreversibly with singlet oxygen. The DPBF optical density at λ=414 nm decreases directly proportional to the fraction of DPBF reacting with singlet oxygen. Briefly, 10 μL of 4 mM DPBF in ethanol were added to 500 μL aliquots of the test samples. Then, each mixture was irradiated with a laser (λ=670 nm; energy density of 0.75 J/cm2). Absorption of the samples was measured at 414 nm before and after irradiation, and the change in optical densities before and after irradiation was used to quantify singlet oxygen production.
Cell Culture. Cancerous (4T1 [murine] and MDA-MB-231 [human]) and noncancerous (NOR-10 [murine] and MCF-10A [human]) cell lines were purchased from American Type Culture Collection (ATCC, Manassas, Va.). All cells were cultured in DMEM, and supplemented with 10% (v:v) fetal bovine serum and 1% (v:v) antibiotic solution (100 IU/mL penicillin and 100 mg/mL streptomycin). All cells were maintained at 37° C. in 5% CO2 and in a humidified atmosphere.
Cell Uptake. ApoHb-Al—PC-Hp uptake by cells was monitored over time by measuring cell fluorescence during the incubation period. Briefly, 4T1, MDA-MB-231, NOR-10, and MCF-10A cells were cultured for 24 hours at a density of 2×103 cells/well. Later, the culture medium was replaced with 200 μL of apoHb-Al—PC-Hp in culture medium at a concentration of 1 μM equivalent concentration of Al-PC and incubated at 37° C. over time. At each time point, the culture medium with apoHb-Al—PC-Hp was removed and stored, the cells were washed twice with PBS, and fresh culture medium was loaded into the well. Next, the fluorescence was read at an excitation of λ=350 nm and emission of λ=680 nm, and the fresh culture media was switched back for culture medium containing apoHb-Al—PC-Hp.
PDT. For PDT, 4T1, MDA-MB-231, NOR-10, and MCF-10A cells were cultured for 24 hours at a density of 5×103 on coverslips. Next, cells were exposed to Al-PC, apoHb-Al-PC-Hp, and apoHb-Hp in culture medium at different concentrations. Later, the cells were irradiated with a laser (λ=670 nm) at energy densities between 0.0 J/cm2 and 4 J/cm2 for 15 minutes in the dark.
Cell Viability. Cell viability was assessed using the MTT assay, which is converted by the mitochondria of viable cells to an insoluble purple precipitate. Briefly, after receiving their treatment, cells were washed with PBS twice and then incubated with 0.5 mg/mL of MTT in culture medium for 2 hours. Then, the MTT solution was washed away, and the purple precipitate was extracted from the cells with 150 μL of dimethyl sulfoxide (DMSO). The absorption was then measured at λ=595 nm using a spectrophotometer (Spectramax; Molecular Devices).
Detection of Fragmented DNA. DNA fragmentation was measured using propidium iodide (PI), which binds to DNA and allows for cell DNA content to be measured via by flow cytometry. Briefly, 4T1, MDA-MB-231, NOR-10, and MCF-10A cells were cultured at a density of 5×105 cells/well for 24 hours. Then, the culture medium was removed, cells were washed twice with PBS, and replaced with medium containing apoHb-Al—PC-Hp at a equivalent concentration of 0.165 μM of Al-PC for 30 min and exposed to laser irradiation (λ=670 nm) applied at 0, 0.5, or 1 J/cm2. After, cells were washed twice with PBS, and the cells were cultured for 24 hours in fresh culture media. The next day, all cells were harvested, centrifuged, and resuspended in PBS. The cell suspension was incubated with PI at 20 μg/mL for 20 minutes in the dark and samples were measured using a FACS scan flow cytometer (BD Biosciences, San Jose, Calif.).
Cell Death Analysis. Cell death by apoptosis or necrosis was analyzed after apoHb-Al—PC-Hp treatment using acridine orange/ethidium bromide double staining. The 4T1, MDA-MB-231, NOR-10, and MCF-10A cells were cultured at a density of 1×104 cells/well for 24 hours. Then, the culture medium was removed, cells were washed twice with PBS, and replaced with medium containing apoHb-Al—PC-Hp at an equivalent concentration of 0.165 μM of Al-PC for 30 min and exposed to laser irradiation (λ=670 nm) applied at 0, 0.5, or 1 J/cm2. After, cells were washed twice with PBS, and the cells were cultured for 24 hours in fresh culture media. The next day, all cells were harvested, centrifuged, and resuspended in PBS. The cell suspension was incubated with 50 μg/mL acridine orange and 50 μg/mL ethidium bromide in the dark. Lastly, the cell suspension was evaluated via fluorescence microscopy to measure the percentage of cells experiencing apoptosis or necrosis.
Results and Discussion
Analysis of Al-PC binding to apoHb. Al-PC in EtOH was prepared as described above and added to PBS or apoHb in PBS. The results of this analysis are shown in
As shown in
In this study, EtOH was used to dissolve Al-PC and maintain it in its monomeric form. However, EtOH can denature and precipitate proteins, thus, the maximum volume ratio of EtOH:apoHb solution was determined which maximized binding of Al-PC to apoHb but prevented protein precipitation. The results are shown in
Based on
After Al-PC bound to apoHb, fresh apoHb-Al-PC was dialyzed against PBS at 4° C. over the course of three days with buffer exchanges at 24 hour intervals. This process was expected to gradually remove EtOH from the product. Unfortunately, the product after dialysis had visible precipitates. Therefore, the resulting solution was sterile filtered through a 0.2 μm syringe filter and the UV-visible spectra was measured (
Synthesis of apoHb-Al-PC-Hp. Given apoHb-Al-PC's instability in aqueous solution, Hp was added at an equimolar ratio to apoHb-Al-PC to stabilize it. Hp is known to bind to apoHb, stabilizing the apoprotein, and to prevent heme release from Hb. Thus, it was hypothesized that formation of the apoHb-Al-PC-Hp complex (APH) would yield a stable protein species. Preliminary results demonstrated that APH was more stable than apoHb-Al-PC, retaining Al-PC absorbance after employing the same dialysis protocol that was used with apoHb-Al-PC. Based on these findings, an Al-PC binding protocol was developed to synthesize APH at large scales as described previously (
Even though, Al-PC concentration and EtOH volume addition were maximized, the results from
From the absorbance and fluorescence spectra of APH (
Assuming that all the added Al-PC bound to apoHb, the total amount of Al-PC added corresponded to an occupation of ˜65% of the active heme-binding sites of apoHb. Interestingly, when considering the total protein content of apoHb and not just the active heme-binding sites, ˜50% of apoHb bound to Al-PC. This may indicate that Al-PC preferentially bound to either the α or β chains of apoHb and did not depend on interactions the proximal histidine in apoHb. Such asymmetric binding could be explained by the higher heme affinity of α chains which may indicate they were the only chains capable of Al-PC incorporation. Moreover, it has been shown that the α chain has a lower helical content than the 0 chain when bound to Hp. A lower helical content (less structure) may allow a chains to more easily bind to heme-like (macrocycles) molecules such as Al-PC. Furthermore, there was no appreciable loss of protein as the 280 nm peak did not decrease during processing (there was a slight increase upon Al-PC addition).
The current repeated Al-PC binding protocol (
Size and stability of APH. The hydrodynamic size, apparent MW and stability of APH were characterized. These results are shown in
Hemoglobin (Hb) tetramers (α2β2) have a diameter of ˜5.5 nm. Upon heme removal from Hb, the apoprotein forms αβ dimers, yielding an expected diameter of ˜2.7 nm. Thus, the DLS measurement of 2.4 nm was similar to the expected size of apoHb. From the DLS measurement, Hp had a hydrodynamic diameter of ˜16 nm with a wide distribution ranging from 12 nm to 26 nm. The wide distribution was expected, since the mixture of Hp2-2 and Hp2-1 polymers used in this study can be composed of different numbers of subunits ranging from 200-900 kDa (αβ Hp dimers). Moreover, after binding apoHb-Al-PC to Hp, a detectable increase of ˜2 nm in diameter was observed which corresponded to apoHb-Al-PC's attachment to Hp. Similar to DLS analysis, the HPLC-SEC chromatograms showed an increase in the apparent MW (i.e. lower elution time) of APH (˜700 kDa) compared to Hp (˜400 kDa). This increase in MW indicated that more than one apoHb was attaching to each Hp binding site which was expected due to the polymeric nature of the Hp species used. Moreover, the final MW of APH was more than an order of magnitude larger than apoHb (MW ˜32 kDa). Thus, in addition to stabilizing the apoHb-Al-PC complex, Hp binding increased the size of the final product (i.e. APH). The larger size of APH can potentially minimize the extravasation of apoHb-Al-PC through blood vessel walls or kidneys which are common pathological traits of cell-free Hb which has a similar size compared to apoHb.
To test the stability of Al-PC in APH, the complex was incubated in PBS and human plasma at 37° C. The decrease in absorbance at 680 nm over time was measured to model the loss of Al-PC activity from the APH complex. The results of this experiment are shown in
Specificity and Retention of Al-PC Binding. Given that heme-binding activity did not correlate to Al-PC binding, the binding reaction of Al-PC was performed with Hp, human serum albumin (HSA), Hb, and reconstituted Hb (rHb; prepared by reinstating heme in apoHb) to assess if Al-PC would bind to proteins without the vacant heme-binding pocket of apoHb. This analysis is shown in
Addition of Al-PC to Hp resulted in a slight increase in the absorbance at 680 nm. This binding event can be attributed to the residual apoHb present in the Hp sample as Hp has been shown previously to not have heme-binding pockets. As shown
To further assess the binding site of Al-PC in apoHb, the APH complex was mixed with heme-albumin at concentrations of 1.5 and 7.5 μM at 37° C. and 4° C. Heme-albumin acted as a carrier for heme that could potentially displace of Al-PC from the heme-binding pocket of apoHb. The change in absorbance of APH at 680 nm was monitored to assess Al-PC loss from APH and the results are shown in
Based on the results of
Assessment of the Al-PC binding site in apoHb via heme-displacement is an important assay for characterizing apoHb complexes. For example, improper analysis of the binding sites has led to a previous study concluding that there were five binding sites per apoHb tetramer (α2β2). In the aforementioned study, apoHb was bound to a Zn-substituted porphyrin moiety. Unfortunately, the previous study employed an incorrect extinction coefficient of apoHb (value ˜1.25× larger) for the apoHb protein, and did not quantify its heme-binding capacity or performed heme-displacement assays. Based on this error, the authors underestimated the quantity of protein in their samples leading to an overestimation of the heme-binding capacity. Furthermore, although Zn-substituted Hb could diminish heme-oxygenase-1 (HO-1) antioxidant activity in tumor cells, the porphyrin provides similar photodynamic characteristics to that of native Hb (absorbance peak between 400-500 nm), and therefore was not an optimal photosensitizer for PDT (the PS should have an absorbance peak at wavelengths above 500 nm to prevent the skin pigment melanin from absorbing light).
Other studies have aimed to improve PDT by binding PS molecules to apomyoglobin. Unfortunately, these studies did not analyze the binding activity of the apoprotein prior to its use or confirm the heme-binding pocket specificity via heme displacement measurements. Moreover, although apomyoglobin-PS led to enhanced photodynamic properties by preventing PS aggregation, the use of apomyoglobin-PS would likely lead to a significant loss of PS through the vascular wall and kidneys given the small MW of myoglobin. Thus, systemic administration of myoglobin-like compounds without addressing its inherently small MW could lead to systemic photosensitivity and toxicity.
In vitro analysis of PDT potential of APH. To analyze the PDT potential of APH, singlet oxygen generation from Al-PC, APH and apoHb-Hp in aqueous solution was assessed and compared to Al-PC in EtOH. Moreover, to determine if APH could be up taken up by normal and human cancer cells, the amount of intracellular Al-PC was tracked after incubating APH with human and murine cancer cell lines. The results from these analyses are shown in
As shown in
With the promising singlet oxygen generating potential of APH and cellular uptake of Al-PC via delivery from APH confirmed, the cytotoxicity of APH, Al-PC, and apoHb-Hp without laser irritation (dark toxicity) as well as the effect of laser irradiation alone were first analyzed. The results from these experiments are shown in
As shown in
Based on the results from
The results from
Conclusion
The potential of PDT for cancer therapy has been known for more than 25 years with recent studies showing its success against various cancers with some therapies now in clinical trials. However, PS solubility and targeting are the main roadblocks limiting clinical application of PDT. In addition to the known benefits of PDT, recent developments of fiber-based interstitial, intravesical and endoscopic light delivery systems have expanded possible applications of PDT. Moreover, new and more potent PSs, such as Al-PCs, already exist, but efficient delivery and targeting mechanisms are needed to minimize systemic photosensitivity, prevent aggregation and increase aqueous solubility. Thus, there is a need to develop systems to deliver PSs to cancer cells, while maintaining photoactivity of the PS. In this example, it was demonstrated how to use apohemoglobin (apoHb) as an efficient carrier of Al-PC by forming the apoHb-Al—PC-Hp (APH) complex. The process developed here is scalable and easily translatable to other PSs or hydrophobic molecules. Further, binding of apoHb-Al-PC to Hp enhances the stability of the complex and provides a potential targeting and fast clearance mechanism via CD163+ macrophage uptake, which are tumor promoting M2 tumor associated macrophages highly expressed in various cancer types (such as rectal, pancreatic, lymphoma, oral squamous cell carcinoma, ovarian, hepatocellular carcinoma, prostate, lung, mesothelioma, brain, and thyroid). Targeted drug delivery could lower the overall systemic toxicity of PS molecules, and the fast clearance could prevent prolonged phototoxicity. Moreover, the large MW and size of APH should prevent extravasation into the tissue space and clearance via the kidneys. Finally, in vitro analysis confirmed APH could generate singlet oxygen and induce light-induced cytotoxicity with minimal dark toxicity. Taken together, this example details a photodynamic agent for PDT. Moreover, the encapsulation protocol described herein may be made continuous and other therapeutic agents may be encapsulated into the apoHb-Hp complex.
The apoHb-Hp-(therapeutic/diagnostic) complex can be made by reacting the apoHb-(therapeutic/diagnostic) conjugate with Hp, or reacting the Hp-(therapeutic/diagnostic) conjugate with apoHb, or conjugating the therapeutic/diagnostic molecule to the apoHb-Hp complex.
Materials and Methods
Apohemoglobin-Haptoglobin Complex Preparation. The apoHb-Hp complex was made by reacting apoHb with Hp. The high binding affinity drives the reaction for complex formation. A Hp solution with a hemoglobin binding capacity (HbBC) of 49.7 mg/mL was mixed with an apoHb solution with 30.3 mg/mL of active apoHb at a 1:2 volume ratio (10 μl of Hp and 20 μl of apoHb in 1 mL of 50 mM phosphate buffer, pH 7.4). The resultant mixture (apoHb-Hp complex+excess apoHb) was separated on a size exclusion chromatography (SEC) column for analysis. Large molecular weight Hp (Hp2-2 and Hp2-1) was mixed with apoHb with a molecular weight of about 31 kDa (dimeric apoHb) and separated on an analytical Acclaim SEC-1000 (4.6×300 mm) column (Thermo Fisher Scientific, Waltham, Mass.) attached to a Dionex UltiMate 3000 system (Thermo Fisher Scientific, Waltham, Mass.). The mobile phase consisted of 50 mM potassium phosphate, pH 7.4. The flow rate and UV-visible spectral detection was controlled using Chromeleon 7 software with detection set to λ=280 nm to detect protein elution at a flow rate 0.35 mL/min. The percent change in the area under the curve between a pure apoHb solution and a mixture of apoHb-Hp with excess apoHb was used to determine the percentage of apoHb that was bound to Hp. This percentage was compared to the mass of pure apoHb loaded to determine the Hp binding capacity of apoHb. This value was compared to the HbBC of the Hp sample.
Apohemoglobin-Haptoglobin-(Therapeutic/Diagnostic) Complex Preparation. To form an apoHb-Hp-(therapeutic/diagnostic) complex, either pure apoHb or apoHb-Hp was mixed with a therapeutic/diagnostic containing solutions. The therapeutic/diagnostic molecule in this case binds to the vacant heme binding pocket of apoHb.
Diagnostic/therapeutic solutions tested here consisted of either Mn-porphyrin IX chloride (Mn-IX) or aluminum phthalocyanine (Al-PC) chloride. The Mn-IX solution was made by dissolving 2.5 mg of Mn-IX in 0.5 mL of 0.1 M NaOH. For Al-PC, 1 mg of Al-PC was dissolved in 1 mL of pure EtOH, then 50 μL of the saturated solution was diluted into 1 mL of pure EtOH (the second dilution ensured that the Al-PC is monomeric in solution). The apoHb-Hp complexes were formed by mixing 20 μL of apoHb (30.3 mg/mL of active apoHb) and 10 μL of Hp (HbBC of 49.7 mg/mL) in PB (50 mM, pH 7.4) for the Mn-IX trials, and 10 μL of apoHb (30.3 mg/mL of active apoHb) and 10 μL of Hp (HbBC of 49.7 mg/mL) in PB (50 mM, pH 7.4) for the Al-PC trials. From the stock diagnostic/therapeutic solutions, either 2 μl of the stock Mn-IX or 100 μl of the Al-PC solutions was added to the protein samples. The resultant mixtures had their absorbance spectras measured via UV-visible spectroscopy and were separated via size exclusion chromatography (SEC) using an Acclaim SEC-1000 (4.6×300 mm) column (Thermo Fisher Scientific, Waltham, Mass.) attached to a Dionex UltiMate 3000 system (Thermo Fisher Scientific, Waltham, Mass.). The mobile phase consisted of 50 mM potassium phosphate, pH 7.4. The flow rate and UV-visible spectral detection was controlled using Chromeleon 7 software with detection set to λ=280 nm (protein), λ=405 nm (residual heme of apoHb and Hp), λ=486 nm (Soret peak of Mn-IX-apoHb), and λ=680 nm (peak of Al-PC-apoHb) at a flow rate 0.35 mL/min.
Results
ApoHb-Hp complex formation is shown in
Prior to HPLC-SEC, the absorbance spectra of each of the samples tested was measured using UV-visible spectroscopy. The results using the Al-PC solution are shown in
The same absorbance spectral analysis was done with the samples made from the Mn-IX solution. These results are shown in
Using the Mn-IX solution, the HPLC-SEC of the complexes formed are shown in
Using the Al-PC solution, the HPLC-SEC of the complexes formed are shown in
β-thalassemia is a genetic hemoglobin (Hb) disorder that affects millions of people world-wide. It is characterized by defective erythropoiesis and anemia, with patients suffering from low levels of abnormal red blood cells (RBCs). The continuous oxygen deficit leads to life-long blood transfusion regimens, which results in iron accumulation toxicity. Moreover, the abnormal RBCs are prone to hemolytic events that release cell-free Hb, heme, and iron, causing oxidative organ and tissue damage. In this study, β-thalassemic mice were treated with the apohemoglobin-haptoglobin (apoHb-Hp) complex for six weeks to simultaneously scavenge cell-free Hb and free heme. Animal weight and RBC parameters were measured throughout the study. Moreover, total iron levels, transferrin concentration and transferrin saturation were measured at the third and sixth week of treatment. At the end of the experiment, spleen and liver weights were measured and markers of liver function were assessed. Furthermore, the total iron content of the liver and spleen was quantified. ApoHb-Hp treatment lowered hepatosplenomegaly, and lowered markers of liver damage. Moreover, apoHb-Hp treatment lead to improved RBC levels, reduced cell fraction of reticulocytes, and prevented an increase in red-blood cell distribution width. Remarkably, apoHb-Hp treatment reduced circulating iron levels, transferrin saturation, increased overall transferrin levels, and lowered iron accumulation within the liver and spleen. These results indicate that scavenging of cell-free Hb and free heme with apoHb-Hp treatment in beta-thalassemia reduced hepatosplenomegaly, normalized RBC levels, and lowered iron accumulation. Based on these outcomes, a mechanism for iron removal via scavenging of cell-free Hb and heme was proposed. Taken together, this study demonstrated that apoHb-Hp can reduce iron toxicity and normalize RBC levels in β-thalassemic animals via scavenging of cell-free Hb and heme. Thus, apoHb-Hp may be a viable treatment strategy to normalize RBCs and iron levels in β-thalassemic patients.
Methods
Apohemoglobin Preparation. The apoHb used in this study was prepared via tangential flow filtration based on the acidic-ethanol heme-extraction procedure as previously described in the literature The heme-binding capacity of apoHb preparations was approximately 80%, with less than 1% residual heme present.
Haptoglobin Preparation. Human Hp was purified from human Cohn fraction IV (FIV) purchased from Seraplex (Pasadena, Calif.) via tangential flow filtration as previously described in the literature. The final protein solution was composed of a mixture of Hp2-1 and Hp2-2 Hp polymers, with an average MW of 400-500 kDa and >95% purity.
Animal Model and Treatment. Thalassemic mice consisted of C57BL/6 heterozygous for the Hbb β-globin gene deletion (Hbbtd3th/BrjK) (beta-thalassemia, Jackson Laboratory). Animals were treated q.o.d. for six weeks with the apoHb-Hp complex (Hp 22.5 mg/mL, apoHb 7.5 mg/mL, 50 μL, n=8), or vehicle (PBS, equal volume as study group, n=8) via tail vein injection. Animal body weight was monitored at alternate treatment days. Animals were sacrificed after the final dose for analysis.
Hematological Parameters. Blood samples were obtained at baseline and every two weeks by retro-orbital puncture under isoflurane (2% for maintenance, Drägerwerk AG, Lübeck, Germany). Complete blood counts (CBC) were measured on an Hemavet blood analyzer (Drew Scientific, Oxford, Conn.) and confirmed via flow cytometry on selected samples.
Serum Iron Content and Tf Saturation. Serum iron and unsaturated iron-binding capacity (UIBC) were measured in non-hemolyzed mouse serum using an iron and total iron-binding capacity (TIBC) assay according to the manufacturer's instructions (LabCorp, Burlington, N.C.). TIBC and transferrin saturation were calculated from the measured serum iron and UIBC.
Determination of Tissue Iron Content. After the experimental end point was reached, half of the mice (n=4/group) were euthanized and transcardially perfused via the aorta with PBS, and tissue non-heme iron content was determined with a colorimetric method using BPS (4,7-diphenyl-1,10-phenantroline disulfonic acid) as the chromogen. Briefly, 0.2 g of tissues were incubated overnight in a mixture of trichloroacetic (10%) and hydrochloric (4 N) acids, and 100 μl of supernatant reduced with thioglycolic acid (Sigma-Aldrich) and acetic acid-acetate buffer (pH 4.5). The ferrous iron content was determined spectrophotometrically (535 nm) with the addition of BPS and after 1 hr incubation at 37° C. The results are expressed as μg iron/g dry tissue weight.
Histology and Iron Staining. After the experimental end point was reached, the other half of the mice (n=4/group) were transcardially perfused via the aorta with PBS followed by perfusion of a fixative solution (4% paraformaldehyde in PBS). The liver and spleen were harvested and were continued to be fixed in the same fixative solution (4 hrs at 4° C.). Tissues were washed in PBS, and cryoprotected by immersion in sucrose overnight. Tissues were cut and the free-floating sections were stored in cryoprotective solution at −20° C. until processed. Iron was detected using Perl's staining for non-heme ferric iron (Fe (III)) followed by 3,3 diamino-benzidine (DAB, Sigma-Aldrich) in methanol. Iron staining was developed by incubation of tissues with DAB and hydrogen peroxide, and then transferred onto gelatin-coated slides, rinsed in PBS, counterstained with hematoxylin, dehydrated and mounted. Quantification of iron positive cells was performed with an Olympus BX51WI microscope equipped with a high-resolution digital CCD ORCA-285 camera (Hamamatsu, Hamamatsu City, Japan). Images for Perl's stained areas and Hoechst stained areas were prepared using Wasabi Imaging Software (Hamamatsu). The ratio of pixels stained for Perl's Prussian Blue in each region compared to the total cellular area of the image was calculated. Ten images were analyzed, by sections, and the results were pooled to determine the mean and SD. To indicate the colocalization of Perl's Prussian Blue and Hoechst in cells, and positive cells were counted.
Statistical Analysis. Results are presented as Tukey box plots. Some data are presented as absolute values and relative to baseline. A ratio of 1.0 signifies no change from baseline, whereas lower or higher ratios are indicative of changes proportionally lower or higher than baseline. Data analysis between groups and time points were analyzed via two-way analysis of variance (ANOVA), with Tukey post-hoc test when appropriate. Before experiments were initiated, sample sizes were calculated based on α=0.05, and power=0.9 to detect differences between primary end points (serum iron and transferrin saturation). All statistics were performed in GraphPad Prism 7 (GraphPad, San Diego, Calif.). Results were considered statistically significant if P<0.05.
Results
This study was completed in sixteen Hbbth3/+ mice. Eight animals were randomly assigned to each experimental group. The first experimental group was untreated, receiving only PBS (vehicle group). The second group was treated with the apoHb-Hp complex (apoHb-Hp group). All animals were confirmed positive for β-thalassemia via genotyping performed by the vendor and tolerated the experiments without signs of pain or discomfort.
Six-weeks of apoHb-Hp treatment does not show signs of toxicity. The toxicity associated with continuous apoHb-Hp treatment was assessed by measuring the body weight of animals during the experiment.
ApoHb-Hp treatment reduces splenomegaly and hepatomegaly induced by β-thalassemia. The enlargement of the spleen (splenomegaly) and liver (hepatomegaly) are common pathological traits of β-thalassemia.
ApoHb-Hp treatment recovers RBC levels in β-thalassemic mice.
When normalized to baseline, the effects of apoHb-Hp treatment were more pronounced. After six weeks of treatment, the relative RBC count, tHb level and Hct of the apoHb-Hp treated group increased compared to baseline and were significantly higher than the control group. Moreover, the relative tHb level of the apoHb-Hp treated group was significantly higher than baseline and vehicle controls at 4 and 6 weeks of treatment. Finally, the relative tHb levels of the control group decreased over the experimental study, becoming significantly lower than baseline at six weeks. These results indicate that apoHb-Hp treatment improved RBC levels, reducing the severity of anemia in β-thalassemic mice. Although the increase in tHb levels also indicated an improvement over the control, the increase could be an artifact of higher cell-free Hb retention when bound to the apoHb-Hp complex. Moreover, the relative decrease in tHb of the control group is indicative of worsening of anemia in β-thalassemic animals, which was not observed in the apoHb-Hp treated animals.
ApoHb-Hp treatment lowers total iron levels and increases serum iron binding capacity of transferrin.
ApoHb-Hp treatment reduces iron accumulation in the liver and spleen.
Discussion
ApoHb-Hp treatment reduced spleen size compared to untreated mice (vehicle control) who most likely suffered from splenomegaly (i.e. enlarged spleen) as it is a common characteristic in β-thalassemic patients. The enlargement of the spleen is generally a result of the increased accumulation of RBCs and iron. This occurs since one of the spleen's primary functions is to remove aged, damaged, or abnormal RBCs from the circulation. The spleen accomplishes this function via its splenic cords in which young, flexible RBCs pass through the epithelial cells of the splenic cord, while senescent RBCs are trapped and phagocytosed by tissue resident macrophages. When presented with an abnormally large number of senescent RBCs, the spleen can become clogged, preventing all RBC passage, and thus RBCs accumulate, contributing to splenomegaly. Moreover, the accumulation of senescent RBCs in the spleen increases macrophage levels which can further block cellular passage, augment splenomegaly, and contribute to increased rates of splenic hemolysis. Notably, it has also been shown that vaso-occlusion induced by hemolysis can increase capture of RBCs within the spleen. Thus, by reducing the hemolysis induced side-effects of β-thalassemia, apoHb-Hp treatment may reduce RBC capture in the spleen, leading to the observed decrease in the size of the organ.
Reduction of splenomegaly is vital for thalassemic patients as it lowers the rate of RBC turnover, by reducing the rate of RBC destruction. The bone marrow in β-thalassemic patients cannot produce enough RBCs to maintain demand, thus causing other organs, such as the spleen and liver, to create RBCs via extramedullary hematopoiesis. Thus, in addition to facilitating the accumulation of damaged RBCs in the spleen, the organ is also stressed due to the need to produce RBCs. These events can lead to a hyperactive spleen (hypersplenism). In hypersplenism, the spleen destroys RBCs at an even faster rate, exacerbating iron toxicity to the point where chelation therapy cannot mitigate it. Treatment at severe stages of hypersplenism primarily consists of splenectomy, which can make patients highly susceptible to infection and sepsis. As shown in the RBC data, apoHb-Hp treatment lead to increased RBC count, lower reticulocyte counts, and lower RBC distribution width. The increase in RBC count and lower RBC distribution width indicates that the RBCs had an increased lifespan in the circulation. Moreover, the decrease in reticulocyte count suggested a suppression of bone marrow erythropoiesis and extramedullary hemoatopoiesis. In addition to directly improving RBC levels, it is interesting to note that apoHb-Hp treatment reduced iron levels within the spleen, liver, and circulation (serum iron and transferrin saturation). These results further corroborate that the overall rate of hemolysis within β-thalassemic mice was reduced given that CD163 expression (receptor responsible for uptake of the Hb-Hp complex) primarily occurs in the liver and spleen, which are the regions of highest RBC/iron accumulation. Thus, these results indicated that apoHb-Hp could reduce the destruction rate of RBCs in β-thalassemic mice, lowering the demand for RBC production and, therefore, aiding in further reduction of splenomegaly, while also normalizing RBC levels.
Lowering the rate of erythropoiesis in β-thalassemia is important, since a common effect of the severe anemia in β-thalassemic patients is inhibition of the hormonal regulator of iron metabolism, hepcidin. Hepcidin binds to ferroportin and induces its intracellular degradation. This mechanism protects tissues from iron overload as it prevents iron efflux from iron-releasing cells such as macrophages and duodenal enterocytes (responsible for absorption of iron in the intestine). In normal physiological states, high circulatory levels of iron and inflammation stimulate hepcidin expression to prevent iron overload in tissues by restricting iron to macrophages and duodenal enterocytes. However, during thalassemia, the continuous expression of erythropoietin blocks hepcidin expression, leading to high ferroportin levels and subsequent iron overload due to high levels of iron absorption. Since apoHb-Hp treatment normalized RBC levels and reduced the erythropoietic drive, the treatment likely normalized hepcidin function, leading to regularization of iron levels in the liver and spleen, and the observed reduction in transferrin iron saturation.
In addition to splenomegaly, β-thalassemic patients commonly suffer from hepatomegaly (enlarged liver). Under normal conditions, the liver stores a majority of iron, but the excess iron pool in β-thalassemic patients is associated with cellular toxicity and leads to enlargement of the liver. Furthermore, similar to splenomegaly, extramedullary hematopoiesis can also occur in the liver, further increasing liver size. Moreover, the chronic hemolytic environment associated with β-thalassemia can lead to excess heme uptake within the liver, resulting in liver damage and congestion due to inflammation and oxidative stress. This causes chronic liver injury and fibrosis in β-thalassemic patients. One of the consequences from extensive liver injury and fibrosis is portal hypertension, which leads to further splenomegaly and increases the risk of hypersplenism. Markers of dysregulated liver function include AST which, when elevated, is indicative of hemolysis or defective erythropoiesis, which is consistent with thalassemia. In this study, apoHb-Hp treatment reduced liver size and reduced the levels of liver damage markers (AST and ALP) indicating that treatment was capable of reducing hemolysis-mediated damage to the organ.
As mentioned previously, the primary cause of splenomegaly in β-thalassemic patients is the accumulation of large quantities of abnormal RBCs in the spleen, which can congest the splenic cords. In β-thalassemia, these abnormalities in RBCs are not only derived from the genetic deficiency in β-globin production, but the hemolytic species (i.e. Hb, heme, and iron) can also lower the lifespan of RBCs. This occurs due to exposure to high levels of ROS, resulting oxidative stress of the RBC membrane, leading to premature macrophage capture. Moreover, liver disease can also lead to abnormal RBCs as the liver controls lipid metabolism. Thus, the damaged liver and excess hemolytic species (i.e. Hb, heme, and iron) can lead to abnormal RBCs, further increasing congestive splenomegaly and anemia.
Based on the results shown in this study and the previously described mechanism of β-thalassemia toxicity, a mechanism of action for apoHb-Hp treatment was proposed and is illustrated in
Consistent with the data presented here and the proposed mechanism of action for apoHb-Hp, previous studies have demonstrated that Hp and Hpx double knock-out mice suffer from splenomegaly and severe liver inflammation and fibrosis. In these prior studies, splenomegaly was primarily caused by the accumulation of RBCs, which was attributed to free heme, leading to vascular alterations that caused adhesion of RBCs to the endothelium. Additionally, Hpx treatment has been shown to reduce liver damage and endothelial function in β-thalassemia and sickle-cell disease. However, Hpx treatment alone did not improve RBC levels in β-thalassemic mice, unlike the apoHb-hp treatment implemented in our study. Moreover, unlike the results shown in our study, one month of Hpx treatment alone increased iron levels in the liver in β-thalassemic mice. While this increase in liver weight would be favorable, as it suggests that the excess-heme was directed to cells specialized in the detoxification of heme, the study demonstrated that Hpx treatment alone could not prevent the morbidities of β-thalassemia.
Conclusion
This study postulated the use of a specific protein complex, apoHb-Hp, to alleviate the toxicities observed in β-thalassemic mice over a six-week treatment period. Treatment with apoHb-Hp did not show any obvious signs of toxicity as indicated by the steady body weight of the animals throughout the study. Furthermore, our data conclusively show a positive outcome in reducing the toxicity that is normally observed in the liver and spleen in patients with β-thalassemia. Moreover, RBC levels were shown to improve compared to untreated mice, which was attributed to normalization of the rate of RBC destruction. Taken together, these data show that the apoHb-Hp complex promotes a positive outcome for animals suffering from β-thalassemia.
The compositions, systems, and methods of the appended claims are not limited in scope by the specific compositions, systems, and methods described herein, which are intended as illustrations of a few aspects of the claims. Any compositions, systems, and methods that are functionally equivalent are intended to fall within the scope of the claims. Various modifications of the compositions, systems, and methods in addition to those shown and described herein are intended to fall within the scope of the appended claims. Further, while only certain representative compositions, systems, and method steps disclosed herein are specifically described, other combinations of the compositions, systems, and method steps also are intended to fall within the scope of the appended claims, even if not specifically recited. Thus, a combination of steps, elements, components, or constituents may be explicitly mentioned herein or less, however, other combinations of steps, elements, components, and constituents are included, even though not explicitly stated.
The term “comprising” and variations thereof as used herein is used synonymously with the term “including” and variations thereof and are open, non-limiting terms. Although the terms “comprising” and “including” have been used herein to describe various embodiments, the terms “consisting essentially of” and “consisting of” can be used in place of “comprising” and “including” to provide for more specific embodiments of the invention and are also disclosed. Other than where noted, all numbers expressing geometries, dimensions, and so forth used in the specification and claims are to be understood at the very least, and not as an attempt to limit the application of the doctrine of equivalents to the scope of the claims, to be construed in light of the number of significant digits and ordinary rounding approaches.
Unless defined otherwise, all technical and scientific terms used herein have the same meanings as commonly understood by one of skill in the art to which the disclosed invention belongs. Publications cited herein and the materials for which they are cited are specifically incorporated by reference.
This application claims benefit of priority of U.S. Provisional Application No. 62/850,315, filed May 20, 2019, U.S. Provisional Application No. 62/850,329, filed May 20, 2019, and U.S. Provisional Application No. 62/994,736, filed Mar. 25, 2020, each of which is hereby incorporated herein by reference.
This invention was made with Government Support under Grant Nos. R56HL123015, R01HL126945, R01EB021926, and R01HL138116 awarded by the National Institutes of Health. The Government has certain rights in the invention.
| Filing Document | Filing Date | Country | Kind |
|---|---|---|---|
| PCT/US2020/033836 | 5/20/2020 | WO | 00 |
| Number | Date | Country | |
|---|---|---|---|
| 62850315 | May 2019 | US | |
| 62850329 | May 2019 | US | |
| 62994736 | Mar 2020 | US |
