Patent Application
20250152633
The present invention relates generally to the fields of cell biology and cellular therapeutics. More particularly, it concerns methods of rejuvenating senescent cells as well as cells made by those methods and their use in therapy.
Bone marrow-derived mesenchymal stem cells (MSCs) are promising candidates for cell-based therapies for many diseases. MSCs are readily available in adult patients, do not require genetic modification for multipotency, and secrete bioactive factors for immunomodulation and angiogenesis (Yong et al., 2018; Clayton et al., 2017). However, current MSC therapies have several key limitations, including suboptimal regenerative phenotype (Murphy et al., 2013), limited potential for in vitro expansion and poor efficacy in aged or diabetic patients (Li et al., 2016). A major key limiter of the regenerative function of MSCs is cellular senescence, which occurs more rapidly in MSCs from aged patients and during cell culture expansion (Zhou et al., 2020). Senescence limits the efficacy of autologous MSC therapies in patients who are elderly (Kuang et al., 2015; Lin et al., 2019) or have co-morbidities like diabetes (Loomans et al., 2004; Vecellio et al., 2014).
Studies have attempted to use MSCs derived from young, allogenic donors to bypass the hurdles poised by senescence in autologous therapies, but the long-term ability of allogenic MSCs to enhance tissue regeneration is limited by an immune response brought on by the transition of the cells from an immunoprivileged to immunogenic state following implantation in the patient (Huang et al., 2010; Isakova et al., 2014). During culture expansion, MSCs experience a loss of differentiation potential and self-renewal capacity (Bonab et al., 2006), shortening of telomeres and chromosomal instability (Drela et al., 2019), metabolic dysfunction (Yuang et al., 2020), and loss of immunomodulatory functions (Fafián-Labora et al., 2019). Thus, senescence limits the usefulness of MSCs from older donors and those with poor health and reduces the in vitro expansion of MSCs, limiting the number of cells that can be generated for therapeutic use.
While the functional characteristics of MSC senescence are well described (Zhou et al., 2020; Turinetto et al., 2016; Chen et al., 2019), the molecular mechanisms that contribute to the development of senescence are less well defined and often multifactorial. Senescent MSCs have limited proliferation capability (Bruder et al., 1997), diminished multipotency (Siegel et al., 2013), and reduced regenerative properties (Yuan et al., 2020). Reactive oxygen species (ROS) byproducts are produced as MSCs age and replicate, causing stress-induced senescence, in which defective antioxidant mechanisms allow for ROS to build up in the mitochondria and lead to metabolic dysfunction and loss of proliferative capability (Kornienko et al., 2019; Denu & Hematti, 2016; Vono et al., 2017). Oxidative stress and other stress stimuli can cause DNA damage, and dysregulated DNA damage repair pathways in aged MSCs can lead to apoptosis or the induction of senescence (Kosar et al., 2011; Banimohamad-Shotorbani et al., 2020). The secretion of soluble factors is also a major aspect in MSC tissue repair and renewal of vasculature (Ahmadi & Rezaie, 2021). As cells become senescent, they adopt a state known as a senescence-associated secretory phenotype (SASP) and cause widespread ECM remodeling and tissue damage via the activation of inflammatory cascades (Lunyak et al., 2017).
Studies have shown that targeting these intrinsic mechanisms of senescence may reverse its progression, providing hope for autologous MSC therapies (Yuang et al., 2020; Stolzing et al., 2008; Meng et al., 2020; Khanh et al., 2020). However, improved methods for reducing cellular senescence in MSC are needed.
Provided herein are biomechanical conditioning treatments for rejuvenating MSCs derived from elderly or diseased patients and MSCs undergoing extensive expansion in cell culture. The enhanced properties of the treated cells provide a significant therapeutic advantage over non-treated, senescent MSCs.
Provided herein are methods of preparing therapeutic cell populations, the methods comprising:
The brachial waveform mechanical stretch in step (b) and/or step (d) may be applied to the cells for 1 day, 2 days, 3 days, 4 days, 5 days, 6 days, 7 days, 8 days, 9 days, or 10 days. The brachial waveform mechanical stretch in step (b) and/or step (d) may be applied to the cells for about 1 hour, about 2 hours, about 3 hours, about 4 hours, about 5 hours, about 6 hours, about 7 hours, or about 8 hours per day. In one aspect, the brachial waveform mechanical stretch in step (b) and/or step (d) is applied to the cells for about 4 hours per day for 7 days.
The brachial waveform in step (b) and/or step (d) may have a frequency of 0.01 Hz-1.00 Hz. The brachial waveform in step (b) and/or step (d) may have a frequency of 0.1 Hz, 0.2 Hz, 0.3 Hz, 0.4 Hz, 0.5 Hz, 0.6 Hz, 0.7 Hz, 0.8 Hz, or 0.9 Hz.
The brachial waveform in step (b) and/or step (d) may have a magnitude of strain of 0.1% to 17.5%. The brachial waveform in step (b) and/or step (d) may have a magnitude of strain of 0.1%, 1%, 2.5%, 5%, 7.5%, 10%, 12.5%, 15%, or 17.5%.
In one aspect, the brachial waveform in step (b) and/or step (d) has a magnitude of 7.5% strain and a frequency of 0.1 Hz.
In some aspects, step (b) of the methods may comprise applying a brachial waveform of mechanical stretch to the starting population of cells for about 4 hours per day for about 7 days; step (c) may comprise expanding the second population of cells under static conditions for at least about 1 week, 2 weeks, 3 weeks, 4 weeks, 5 weeks, 6 weeks, 7 weeks, or 8 weeks; and step (d) may comprise applying a brachial waveform of mechanical stretch to the expanded second population of cells in the presence of at least one pharmacological agent that induces differentiation of the cells for about 4 hours per day for about 7 days.
The at least one pharmacological agent that induces differentiation of the cells may be an agent that inhibits signaling of at least one ErbB family protein. The agent that inhibits signaling of at least one ErbB family protein may inhibit EGFR/ErbB1 signaling, HER2/ErbB2 signaling, HER4/ErbB4 signaling, or EGFR/PKC signaling. The agent that inhibits signaling of at least one ErbB family protein may enhance Smad2/3 and/or Hippo pathway activation. The agent that inhibits signaling of at least one ErbB family protein may be a kinase inhibitor, such as, for example, an EGFR/Erb-2/4 inhibitor or a PKCβII/EGFR inhibitor.
In some aspects, step (b) may comprise applying a brachial waveform of mechanical stretch to the starting population of cells for about 4 hours per day for about 7 days; step (c) may comprise expanding the second population of cells under static conditions for at least about 1 week, 2 weeks, 3 weeks, 4 weeks, 5 weeks, 6 weeks, 7 weeks, or 8 weeks; and step (d) may comprise applying a brachial waveform of mechanical stretch to the expanded second population of cells in the presence of at least one pharmacological agent that inhibits signaling of at least one ErbB family protein for about 4 hours per day for about 7 days.
The starting population of cells may be or may have been obtained from a donor at least 60 years of age. The starting population of cells may be or may have been obtained from a diabetic donor. The starting population of cells may have previously undergone in vitro expansion. The starting population of cells may be senescent.
The cells may be mesenchymal stem cells. The cells may be senescent mesenchymal stem cells. The cells may be T cells. The cells may be CAR T cells. The cells may be a therapeutic cell type or cell type used in organotypic models of disease or physiology such as human progenitor cells, endothelial cells, vascular smooth muscles, fibroblasts, cardiomyocytes, skin cells, liver cells, or epithelial cells.
The methods may reverse the senescence of the starting cell population. The population of cells produced by step (d) may comprise fewer senescent cells as compared to the starting population of cells. The method may prevent senescence in the population of therapeutic cells.
The population of therapeutic cells may have increased multipotency as compared to the starting population of cells. The population of therapeutic cells may have increased multipotency as compared to a cell population generated by culturing the starting population of cells under static conditions. The population of therapeutic cells may have increased proliferative capacity as compared to the starting population of cells. The population of therapeutic cells may have increased proliferative capacity as compared to a cell population generated by culturing the starting population of cells under static conditions. The population of therapeutic cells may have improved regenerative properties as compared to the starting population of cells. The population of therapeutic cells may have improved regenerative properties as compared to a cell population generated by culturing the starting population of cells under static conditions. The population of therapeutic cells may comprise fewer senescent cells as compared to the starting population of cells. The population of therapeutic cells may comprise fewer senescent cells as compared to a cell population generated by culturing the starting population of cells under static conditions.
The second population of cells may have an enhanced proliferative function as compared to the starting population of cells. The second population of cells may have an enhanced proliferative capacity as compared to a cell population generated by culturing the starting population of cells under static conditions. The second population of cells may comprise fewer senescent cells as compared to the starting population of cells. The second population of cells may comprise fewer senescent cells as compared to a cell population generated by culturing the starting population of cells under static conditions. The second population of cells may comprise decreased DNA damage as compared to the starting population of cells. The second population of cells may comprise decreased DNA damage as compared to a cell population generated by culturing the starting population of cells under static conditions.
The methods may further comprise step (e) encapsulating the population of therapeutic cells in a gel delivery vehicle. The gel delivery vehicle may be an alginate-RGD-collagen gel delivery vehicle.
The methods may further comprise step (e) culturing the population of therapeutic cells on a tissue engineering scaffold to generate a tissue-engineered construct.
Provided herein are populations of therapeutic cells produced by the methods described herein. The populations of therapeutic cells produced by the methods described herein may be used in the manufacture of a medicament for the treatment of a disease or an injury in a patient.
Provided herein are compositions comprising the populations of therapeutic cells described herein. The cells may be encapsulated in a gel delivery vehicle. The cells may be encapsulated in an alginate-RGD-collagen gel delivery vehicle. The cells may be comprised in a tissue-engineered construct. The tissue engineering scaffold may be a vascular graft, an arteriovenous graft, an artery graft, a vein graft, or a coronary graft. The compositions may be for use in the treatment of a patient with a disease or an injury.
Provided herein are methods of treating a patient in need thereof comprising administering to the patient a therapeutically effective amount of the compositions comprising the populations of therapeutic cells described herein. The patient may have a cardiovascular disease, may be receiving vascular grafts, may have ischemia, may have a healing wound, may have peripheral ischemia, or may have peripheral vascular disease.
Administering may comprise injecting or implanting the cells into the patient. The composition or the cells may be implanted into the patient at the site of diseased tissue.
The composition or the cells may be implanted into the patient at the site of vascular dysfunction. The patient may have peripheral artery disease or ischemia. The methods may induce therapeutic angiogenesis.
The starting population of cells may be mesenchymal stem cells harvested from the bone marrow or adipose tissue of the patient. The population of therapeutic cells may be autologous to the patient.
Other objects, features and advantages of the present invention will become apparent from the following detailed description. It should be understood, however, that the detailed description and the specific examples, while indicating preferred embodiments of the invention, are given by way of illustration only, since various changes and modifications within the spirit and scope of the invention will become apparent to those skilled in the art from this detailed description.
The patent or application file contains at least one drawing executed in color. Copies of this patent or patent application publication with color drawing(s) will be provided by the Office upon request and payment of the necessary fee.
The following drawings form part of the present specification and are included to further demonstrate certain aspects of the present invention. The invention may be better understood by reference to one or more of these drawings in combination with the detailed description of specific embodiments presented herein.
Mesenchymal stem cells are a promising cell type for autologous regenerative therapies, but clinical trials have largely yielded inconsistent and disappointing results. A major reason for these limitations is cellular senescence, which is a state in which a cell loses its capacity to replicate and/or loses its therapeutic and functional properties due to cellular aging. Cellular senescence is caused by poor donor health (Loomans et al., 2004; Vecellio et al., 2014), advanced age (Kuang et al., 2015; Lin et al., 2019), and the extensive in vitro expansion required to generate an adequate number of cells for therapeutic applications (Zhou et al., 2020). Senescence also causes MSCs to lose their multipotency, genetic stability, and immunomodulatory functions, drastically limiting their efficacy in autologous therapies. Senescent MSCs are known to secrete pro-inflammatory factors that worsen tissue regeneration and remodeling and spread the senescence to other cells in their microenvironment.
Mechanically conditioning MSCs derived from aged donors enhances their short and long-term proliferative function, improves their multipotency, and protects the cells from advancing senescence, resulting in a lower population of senescent cell phenotypes following the treatment. Brachial mechanical loading in co-treatment with an EGFR/ErbB-2/4 (E/E) synergistically enhances the angiogenic properties of MSCs (Lee et al., 2021a). However, brachial loading alone is superior in enhancing aged MSC proliferative function, expansion capacity, and multipotent properties. In this approach, brachial loading alone is first used to improve the efficiency of MSC culture expansion. Once a sufficient population of healthy cells are generated, the E/E inhibitor is added as a cotreatment to drive MSC differentiation into vascular phenotypes with improved regenerative properties before delivery to patients with peripheral artery disease and ischemia. The brachial loading as a first step to MSC expansion could also be useful in many other applications, if later followed by differentiation into other phenotypes. By conditioning the cells for a short time each day for one week, they have enhanced expansion capacity including increased proliferation and expansion in the long term. For instance, growing 10,000 MSCs out for 7 passages using prior methods could expand these into around 1 million cells. Using the methods described herein, 10,000 MSCs can be expanded into 9.8 million cells.
Mechanical loading may be universally applied to enhance the expansion of various cell cultures. According to a computational model (Miles et al., 2023), mechanical strains within an optimal range may exert forces on chromatin, potentially leading to the opening of DNA without DNA damage. Notably, from the model of cell stretching, brachial loading appears to have a broader range where it can facilitate DNA opening without causing damage. This empirical evidence combined of these studies with these models suggests that a range of cell types might similarly benefit from such biophysical interventions to augment their culture expansion, diminish senescence, and stimulate DNA damage repair mechanisms. To explore this hypothesis further, high-passage human umbilical vein endothelial cells (HUVECs) at passage 10 were treated with brachial loading. Their proliferation, β-galactosidase expression, and capacity for long-term culture expansion were measured. While brachial loading did not enhance short-term DNA synthesis, it significantly decreased the number of β-galactosidase positive cells and markedly improved the cumulative doubling rate during extended in vitro culture (
Brachial mechanical loading results in the upregulation of several Sirtuin proteins, which are known to be crucial regulators of aging. Several current clinical trials are investigating the anti-aging potential of pharmacological SIRT1 activators such as Resveratrol (Pyo et al., 2020) and Metformin (Chalasani et al., 2018), although results remain inconclusive in human subjects. Sirtuin proteins may alter key pathways of senescence, oxidative stress management and DNA damage repair, to rejuvenate the functionality of aged MSCs. Previous work has shown that SIRT6 functions as a DNA damage sensor and activates DNA damage signaling through the recruitment and phosphorylation of ATM and γ-H2AX histone foci (Onn et al., 2020). The activated ATM in turn phosphorylates Akt to promote cell survival and proliferation (Halaby et al., 2008), which is in agreement with the results of the mechanistic studies provided herein (
Mechanical conditioning of aged MSCs results in the enhanced recognition and repair of DNA double-strand-breaks. Enhanced DNA damage repair was not dependent on Piezo1-mediated mechanosensitive signaling and only partially dependent on import of nuclear proteins through importin-1. In addition, mechanical loading caused enhanced recognition of DNA damage leading its repair over time. Together, these findings suggest a biophysical mechanism in which mechanical loading reveals occult DNA damage that can then be repaired. Mechanical conditioning also resulted in the broad activation of antioxidant proteins such as SOD1, FOX01, FOX03a and FOX04, and DNA damage repair proteins such as KU80 and ATM. Mechanical conditioning also reduced the buildup of reactive oxygen species and resulted in rapid repair of DNA damage. When oxidative signaling was altered through the co-treatment of aged MSCs with the antioxidant N-Acetyl Cysteine (NAC) and brachial loading, a reduction of proliferative enhancement was observed. Conversely, when DNA damage repair signaling was altered through the co-treatment of ATM inhibitor KU55933 with brachial loading, a reduction of senescence-protective effects was observed, resulting in a larger phenotype of β-Galactosidase+ senescent cell phenotypes. Together, these results suggest that mechanical conditioning relies upon altered signaling in both key pathways of senescence to rejuvenate specific aspects of the cells' functionality.
Several studies have targeted the elevated reactive oxygen species (ROS) and impaired antioxidant mechanisms that are characteristic of senescent MSCs, using gene vectors to overexpress the Sirtuin proteins (Meng et al., 2020; Zhang et al., 2018), antioxidant pharmaceutical treatment (Lee et al., 2016; Park et al., 2017), or conditioning with extracellular exosomes (Khanh et al., 2020). However, additional studies have shown that high doses of antioxidant expression can lead to DNA damage and induce premature senescence, and there remains a need to further evaluate anti-aging antioxidant potential (Kornienko et al., 2019). Other studies have shown that over-activation of the DNA damage repair network can transiently halt cell cycle progression in aged MSCs and persistence of DNA damage results in activation of tumor suppressor functions to induce senescence (Zhang et al., 2009; Ciccia & Elledge, 2010). Mechanical conditioning led to upregulated antioxidant expression and enhanced DNA damage repair, suggesting that there is a crucial homeostatic balance of signaling in and between these pathways in the rejuvenation of senescence. Therefore, while highly targeted attempts to improve the antioxidant or DNA damage repair response genetically or pharmacologically may cause unintended consequences, mechanical treatment resulted in the broad enhancement of these pathways in aged MSCs to restore their functionality. Taken together, these findings provide a practical method to enhance the regenerative function of MSCs in aged patients.
While this technique has been tested in the rejuvenation of aged MSCs, it is expected that other cell types would receive similar functional enhancements. Senescence is not unique to MSCs, but also limits cell therapies for other diseases including cancer (e.g., CAR-T, T cells), chronic kidney disease (e.g., epithelial cells, endothelial cells), vascular disease and tissue engineered grafts (e.g., smooth muscle cells, endothelial cells), metabolic disorders, and inflammatory conditions (Huang et al., 2022). While the specific use of the treated cells would vary for each of these therapeutic indications, all are currently limited by the diminished proliferation, expansion capacity, and regenerative potential of the senescent cells.
As used herein, “essentially free,” in terms of a specified component, is used herein to mean that none of the specified component has been purposefully formulated into a composition and/or is present only as a contaminant or in trace amounts. The total amount of the specified component resulting from any unintended contamination of a composition is therefore well below 0.05%, preferably below 0.01%. Most preferred is a composition in which no amount of the specified component can be detected with standard analytical methods.
As used herein the specification, “a” or “an” may mean one or more. As used herein in the claim(s), when used in conjunction with the word “comprising,” the words “a” or “an” may mean one or more than one.
The use of the term “or” in the claims is used to mean “and/or” unless explicitly indicated to refer to alternatives only or the alternatives are mutually exclusive, although the disclosure supports a definition that refers to only alternatives and “and/or.” As used herein “another” may mean at least a second or more.
Throughout this application, the term “about” is used to indicate that a value includes the inherent variation of error for the device, the inherent variation in the method being employed to determine the value, the variation that exists among the study subjects, or a value that is within 10% of a stated value.
The term “cell population” is used herein to refer to a group of cells, typically of a common type. The cell population can be derived from a common progenitor or may comprise more than one cell type. An “enriched” cell population refers to a cell population derived from a starting cell population (e.g., an unfractionated, heterogeneous cell population) that contains a greater percentage of a specific cell type than the percentage of that cell type in the starting population. The cell populations may be enriched for one or more cell types and depleted of one or more cell types.
The term “stem cell” refers herein to a cell that under suitable conditions is capable of differentiating into a diverse range of specialized cell types, while under other suitable conditions is capable of self-renewing and remaining in an essentially undifferentiated pluripotent state. The term “stem cell” also encompasses a pluripotent cell, multipotent cell, precursor cell and progenitor cell. Exemplary human stem cells can be obtained from hematopoietic or mesenchymal stem cells obtained from bone marrow tissue, embryonic stem cells obtained from embryonic tissue, or embryonic germ cells obtained from genital tissue of a fetus. Exemplary pluripotent stem cells can also be produced from somatic cells by reprogramming them to a pluripotent state by the expression of certain transcription factors associated with pluripotency; these cells are called “induced pluripotent stem cells” or “iPSCs.”
The term “pluripotent” refers to the property of a cell to differentiate into all other cell types in an organism, with the exception of extraembryonic, or placental, cells. Pluripotent stem cells are capable of differentiating to cell types of all three germ layers (e.g., ectodermal, mesodermal, and endodermal cell types) even after prolonged culture. A pluripotent stem cell is an embryonic stem cell derived from the inner cell mass of a blastocyst. In other embodiments, the pluripotent stem cell is an induced pluripotent stem cell derived by reprogramming somatic cells.
The term “differentiation” refers to the process by which an unspecialized cell becomes a more specialized type with changes in structural and/or functional properties. The mature cell typically has altered cellular structure and tissue-specific proteins.
As used herein, “undifferentiated” refers to cells that display characteristic markers and morphological characteristics of undifferentiated cells that clearly distinguish them from terminally differentiated cells of embryo or adult origin.
An “isolated” cell has been substantially separated or purified from others cells in an organism or culture. Isolated cells can be, for example, at least 99%, at least 98% pure, at least 95% pure or at least 90% pure.
An “embryonic stem (ES) cell” is an undifferentiated pluripotent cell which is obtained from an embryo in an early stage, such as the inner cell mass at the blastocyst stage, or produced by artificial means (e.g. nuclear transfer) and can give rise to any differentiated cell type in an embryo or an adult, including germ cells (e.g. sperm and eggs).
“Induced pluripotent stem cells (iPSCs)” are cells generated by reprogramming a somatic cell by expressing or inducing expression of a combination of factors (herein referred to as reprogramming factors). iPSCs can be generated using fetal, postnatal, newborn, juvenile, or adult somatic cells. In certain embodiments, factors that can be used to reprogram somatic cells to pluripotent stem cells include, for example, Oct4 (sometimes referred to as Oct 3/4), Sox2, c-Myc, and Klf4, Nanog, and Lin28. In some embodiments, somatic cells are reprogrammed by expressing at least two reprogramming factors, at least three reprogramming factors, or four reprogramming factors to reprogram a somatic cell to a pluripotent stem cell.
A “therapeutically effective amount” used herein refers to the amount of a compound that, when administered to a subject for treatment of a disease or condition, is sufficient to effect such treatment.
A “brachial waveform” as used herein refers to a waveform with 2 or more peaks during one cycle. A brachial waveform may mimic the distension of arteries during the cardiac cycle. For example, a brachial waveform may consist of a strain waveform that has two peaks in one cycle, with the first peak being higher than the second peak.
Peripheral vascular disease (PVD) can result from atherosclerotic occlusion of the blood vessels, particularly in limbs and areas distal from the heart, resulting in diminished blood flow and insufficient oxygen perfusion to tissues in the vicinity of and downstream from the occlusion. PVD is frequently manifested in the iliac blood vessels, femoral and popliteal blood vessels, and subclavian blood vessels, and its effects can be exacerbated by thrombi, emboli, or trauma. It is estimated that approximately 8 to 12 million individuals in the United States, especially among the elderly population and those with diabetes, are afflicted with PVD.
Common symptoms of PVD include cramping in the upper and lower limbs and extremities, numbness, weakness, muscle fatigue, pain in the limbs and extremities, hypothermia in the limbs and extremities, discoloration of the extremities, dry or scaly skin, and hypertension. The most common symptom is claudication or feelings of pain, tightness, and fatigue in muscles downstream of the occluded blood vessel that occurs during some form of exercise such as walking, but self-resolve after a period of rest.
In terms of pathophysiology, the occluded blood vessels cause ischemia of tissues at the site of and distal to the obstruction. This ischemia is generally referred to as peripheral ischemia, meaning that it occurs in locations distal to the heart. The severity of the ischemia is a function of the size and number of obstructions, whether the obstruction is near a muscle or organ, and whether there is sufficient redundant vasculature. In more severe cases, the ischemia results in death of the affected tissues, and can result in amputation of affected limbs, or even death of the patient.
Current methods for treatment of more severe cases of PVD include chemotherapeutic regimens, angioplasty, insertion of stents, reconstructive surgery, bypass grafts, resection of affected tissues, or amputation. Unfortunately, for many patients, such interventions show only limited success, and many patients experience a worsening of the conditions or symptoms.
An emerging strategy for treating ischemia is to use growth factors, gene therapies, or regenerative cell types to stimulate tissue repair via the generation of new blood vessels. (Veith, A. P., et al. (2019). “Therapeutic strategies for enhancing angiogenesis in wound healing.” Advanced Drug Delivery Reviews146: 97-125.) (Carmeliet, P. and M. Baes (2008). “Metabolism and Therapeutic Angiogenesis.” New England Journal of Medicine358 (23): 2511-2512.)
Presently, there is interest in using either stem cells, which can divide and differentiate, or muscles cells from other sources, including smooth and skeletal muscles cells, to assist the in the repair or reversal of tissue damage. Transplantation of stem cells can be utilized as a clinical tool for reconstituting a target tissue, thereby restoring physiologic and anatomic functionality. The application of stem cell technology is wide-ranging, including tissue engineering, gene therapy delivery, and cell therapeutics, i.e., delivery of biotherapeutic agents to a target location via exogenously supplied living cells or cellular components that produce or contain those agents. The identification of stem cells has stimulated research aimed at the selective generation of specific cell types for regenerative medicine.
One obstacle to realization of the therapeutic potential of stem cell technology has been the difficulty of obtaining sufficient numbers of stem cells. A reliable, well-characterized and plentiful supply of substantially homogeneous populations of such cells having the ability to differentiate into an array of skeletal muscle, pericyte, or vascular lineages would be an advantage in a variety of diagnostic and therapeutic applications for skeletal muscle repair, regeneration, and improvement, for the stimulation and/or support of angiogenesis, and for the improvement of blood flow subsequent to a peripheral ischemic event, particularly in PVD patients.
The pharmaceutical compositions provided herein may be administered at sites of peripheral ischemia. The pharmaceutical composition may be administered locally. The pharmaceutical composition may be administered by injection, infusion, a device implanted in a patient, or by implantation of a matrix or scaffold containing the pharmaceutical composition. The pharmaceutical composition may be administered by injection into interstitial spaces so as not to enter directly into circulation. The isolated population of cells may be induced in vitro to differentiate into a skeletal muscle, vascular muscle, pericyte or vascular endothelium lineage prior to administration. The population of cells may also be genetically engineered to produce a gene product that promotes treatment of peripheral vascular disease. Optionally, the composition further comprises an agent selected from the group consisting of an antithrombogenic agent, an immunosuppressive agent, an immunomodulatory agent, a pro-angiogenic, an antiapoptotic agent and mixtures thereof.
Intra-arterial administration (IA) involves delivery of the pharmaceutical compositions into at least one artery. In an IA infusion, the infusion is typically divided into several arteries in the legs, e.g., the left and right common femoral arteries, but is sometimes administered into a single artery. The infusion can be administered for about 1 minute, 1 to 5 minutes, 10 to 20 minutes, or 20 to 30 minutes into each artery in both legs. The infusion can be repeated from time to time to achieve or sustain the predicted benefit. The timing for repeat administration is based on the patient's response as measured by symptoms and hemodynamic measures.
Providing a therapy or “treating” refers to indicia of success in the treatment or amelioration of an injury, disease or condition, including any objective or subjective parameter such as abatement, remission, diminishing of symptoms of making the injury, disease or condition more tolerable to the patient, slowing the rate of degeneration or decline, making the final point of degeneration less debilitating, or improving a patient's physical or mental well-being. Those in need of treatment include those already with the disease or condition as well as those prone to have the disease or condition or those in whom the disease or condition is to be prevented. Preferred subjects for treatment include animals, most preferably mammalian species, such as humans and domestic animals such as dogs, cats, and the like, subject to the disease and other conditions. A “patient” refers to a subject, preferably mammalian (including human). Where the specification indicates that a number of cells are to be administered, a person of ordinary skill in the art will understand that these are approximate values.
In some embodiments, the dosage of the composition encompassing a therapeutically effective amount of mesenchymal stem cells ranges from 1.0×103-1.0×108 cells/kg (weight). In some embodiments, the dosage ranges from 1.0×104-1.0×107 cells/kg (weight). In some embodiments, about 1.0×103 cells/kg, about 1.0×104 cells/kg, about 1.0×105 cells/kg, about 1.0×106 cells/kg, about 1.0×107 cells/kg or about 1.0×108 cells/kg are administered.
The dosage of the composition may vary depending on patient's weight, age, sex and symptoms, the dosage form of the composition to be administered, a method of administering the composition, and so on. The frequency of administration may range from one to several times. There may be one or more administration sites. The dosage per kg for non-human animals may be the same as that for human, or can be converted from the above-described dosage, for example, based on the volume ratio (for example, average value) between the diseased tissue of the human and animal subjects. Animals to be treated according to the present invention include human and other desired mammals, specific examples of which include humans, monkeys, mice, rats, rabbits, sheep, horses, cats, cows and dogs.
In accordance with the present invention, the diseases or conditions can be treated or prevented by intravenous administration of the mesenchymal stem cells described herein. In some embodiments, about 20 million, about 40 million, about 60 million, about 80 million, about 100 million, about 120 million, about 140 million, about 160 million, about 180 million, about 200 million, about 220 million, about 240 million, about 260 million, about 280 million, about 300 million, about 320 million, about 340 million, about 360 million, about 380 million, about 400 million, about 420 million, about 440 million, about 460 million, about 480 million, about 500 million, about 520 million, about 540 million, about 560 million, about 580 million, about 600 million, about 620 million, about 640 million, about 660 million, about 680 million, about 700 million, about 720 million, about 740 million, about 760 million, about 780 million, about 800 million, about 820 million, about 840 million, about 860 million, about 880 million, about 900 million, about 920 million, about 940 million, about 960 million, or about 980 million cells are injected intravenously. In some embodiments, about 1 billion, about 2 billion, about 3 billion, about 4 billion or about 5 billion cells or more are injected intravenously. In some embodiments, the number of cells ranges from between about 20 million to about 4 billion cells, between about 40 million to about 1 billion cells, between about 60 million to about 750 million cells, between about 80 million to about 400 million cells, between about 100 million to about 350 million cells, and between about 175 million to about 250 million cells.
In some embodiments, a single intravenous administration is sufficient, while in other embodiments, multiple intravenous administrations are performed, such as 2, 3, 4, 5, 6, 7, 8, 9 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, or 20 intravenous administrations of the mesenchymal stem cells. The treatment interval(s) can be spaced such that an administration follows a prior administration by one day, 2 days, 3 days, 4 days, 5 days, 6 days, one week, 1½-2 weeks, 3 weeks, one month, or 2-3 months, 6 months, one year, or two years or longer. In some embodiments, the treatment interval is spaced in accordance with the progression of the patient's improvement or response to treatment. For example, in some embodiments, a first treatment is administered followed by a second treatment one week later, followed by a third treatment one month later, followed by a fourth treatment 6 months later.
The therapeutic methods of the present invention can be conducted alone or in combination with other standard or advanced methods or pharmaceutical treatments.
The therapeutic composition of mesenchymal stem cells for use in the methods of the present invention can comprise pharmaceutically acceptable carriers and/or additives. Examples thereof include sterilized water, physiological saline, a standard buffer (e.g., phosphoric acid, citric acid, or other organic acids), a stabilizer, salt, an antioxidant (e.g., ascorbic acid), a surfactant, a suspending agent, an isotonic agent, or a preservative. As used herein, the term “base” refers to a base solution in which the mesenchymal stem cells in the cell therapeutic composition are suspended. In some embodiments, physiological saline, phosphate buffered saline or Hartrnan-D (Choongwae Pharma Corp.) is used as the base solution.
In some embodiments, the cell therapeutic composition is prepared in a dosage form suitable for injection. In some embodiments, the mesenchymal stem cells are dissolved (suspended) in a pharmaceutically acceptable aqueous solution, or frozen in a solution state. The kit of the present invention may further comprise a desired pharmaceutically acceptable carrier that can be used to suspend or dilute the mesenchymal stem cells. Examples of such a carrier include distilled water, physiological saline, PBS and the like.
The composition for use in the present invention can contain a pharmaceutically acceptable carrier or excipient, or any necessary stabilizer or adsorption-preventing agent to provide a pharmaceutical preparation that is suitable for administration to humans or animals. The composition of the present invention can be formulated in the form of an injectable solution (e.g., injection solutions for subcutaneous, intradermal, intramuscular, intravenous and intraperitoneal injection). In some embodiments, upon the injection of the composition of mesenchymal stem cells, an analgesic agent, which can relieve pains, may be used.
The cell therapeutic composition of mesenchymal stem cells for use in the present invention can be filled into a syringe, a device, a cryovial in which cells can be frozen, or a pyrogen-free glass vial comprising rubber stoppers and aluminum caps, which contains liquid drugs.
The cell therapeutic composition of mesenchymal stem cells for use in the present invention can, if necessary, contain at least one selected from among suspending agent, solubilizing agents, stabilizers, isotonic agents, preservatives, adsorption-preventing agents, surfactants, diluents, vehicles, pH-adjusting agents, analgesic agents, buffering agents, sulfur-containing reducing agents and antioxidants, depending on the administration mode or formulation thereof.
Examples of the suspending agents may include methylcellulose, Polysorbate 80, hydroxyethylcellulose, gum acacia, gum tragacanth powder, sodium carboxymethylcellulose, polyoxyethylene sorbitan monolaurate, etc. The solubilizing agents include polyoxyethylene hydrogenated castor oil, polysorbate 80, nicotinamide, polyoxyethylene sorbitan monolaurate, Macrogol and castor oil fatty acid ethyl esters. The stabilizers include dextran 40, methylcellulose, gelatin, sodium sulfite, sodium metasulfite, etc. Examples of the isotonic agents are D-mannitol and sorbitol.
Examples of the preservatives include methyl parahydroxybenzoate, ethyl parahydroxybenzoate, sorbic acid, phenol, cresol, and chlorocresol. Examples of the adsorption preventing agents include human serum albumin, lecithin, dextran, ethylene oxide-propylene oxide copolymer, hydroxypropylcellulose, methylcellulose, polyoxyethylene hydrogenated castor oil, and polyethylene glycol.
The sulfur-containing reducing agents include N-acetylcysteine, N-acetylhomocysteine, thioctic acid, thiodiglycol, thioethanolamine, thioglycerol, thiosorbitol, thioglycolic acid and salts thereof, sodium thiosulfate, glutathione, and sulfhydryl-containing compounds such as thioalkanoic acid having 1 to 7 carbon atoms.
The antioxidants include, for example, erythorbic acid, dibutylhydroxytoluene, butylhydroxyanisole, [alpha]-tocopherol, tocopherol acetate, L-ascorbic acid and salts thereof, L-ascorbyl palmitate, L-ascorbyl stearate, sodium bisulfite, sodium sulfite, triamyl gallate, propyl gallate or chelating agents such as disodium ethylenediamine tetraacetate (EDTA), sodium pyrophosphate and sodium metaphosphate. The cryopreservatives include, for example, DMSO, glycerol, etc.
Furthermore, in some embodiments, the cell therapeutic composition of mesenchymal stem cells for use in the methods of the present invention can comprise conventional additives, such as inorganic salts, including sodium chloride, potassium chloride, calcium chloride, sodium phosphate, potassium phosphate and sodium hydrogen carbonate, and organic salts, including sodium citrate, potassium citrate and sodium acetate.
In some embodiments, sucrose or albumin is added to the mesenchymal stem cells to improve stability, prior to cold storage of the cells. In some embodiments, the cells are combined with physiological saline, sucrose, albumin and cryopreservative DMSO prior to freezing and cold storing the cells.
The following examples are included to demonstrate preferred embodiments of the invention. It should be appreciated by those of skill in the art that the techniques disclosed in the examples which follow represent techniques discovered by the inventor to function well in the practice of the invention, and thus can be considered to constitute preferred modes for its practice. However, those of skill in the art should, in light of the present disclosure, appreciate that many changes can be made in the specific embodiments which are disclosed and still obtain a like or similar result without departing from the spirit and scope of the invention.
The inventors have developed a system that enables the application of mechanical stretch in a high throughput format. Using this system, the inventors have demonstrated methods for using specific mechanical loading regimes and pharmacological compounds to enhance the regenerative capacity of MSCs from young donors for treating peripheral ischemia (Lee et al., 2021a). In these studies, maximum angiogenic signal responsiveness was achieved with a mechanical loading regime of 4 hours per day for 7 days of 7.5% maximal strain at 0.1 Hz frequency of loading using a physiological waveform similar to stretch of the brachial artery during the cardiac cycle (brachial loading) (Lee et al., 2013). In addition, 1 μM EGFR/ErbB-2/4 (E/E) inhibitor as a cotreatment with loading led to a dramatic increase in MSCs that expressed the markers for both endothelial cells and pericytes, and the resulting cells had enhanced angiogenic activity in a tube formation assay (Lee et al., 2021a).
To investigate the treatments' effects on gene expression patterns that are correlated with senescence in MSCs, MSCs were obtained from three young, healthy donors, and conditioned with brachial loading and E/E inhibitor. Following the treatments, RNA-seq analysis was performed on the cells, and the expression of several key gene sets related to various mechanisms of senescence in MSCs was evaluated. These gene sets were identified through a thorough investigation of the literature as key mediators of MSC senescence, and the expression pattern of each gene as it pertains to MSC senescence was documented (Zhou et al., 2020; Brandl et al., 2011; Turinetto et al., 2016; Ozkul & Galderisi, 2016; Fridman & Tainsky, 2008; Ki et al., 2014; Cakouros & Gronthos, 2019; Peffers et al., 2016). The analysis found that the treatments reversed the effects implicated by senescence on genes related to age-related senescence, oncogenic-related senescence, stress-related senescence, developmental-related senescence, and inflammation-mediated senescence (
To test if mechanical loading could improve the proliferative function of aged MSCs, MSCs from three donors (ranging from 68-92 years of age) were conditioned with a brachial waveform at a maximal strain of 7.5% for 7 days for 4 hours per day in combination with 1 μM EGFR/ErbB-2/4 (E/E) inhibitor. After the treatments, cells were evaluated for DNA synthesis with a bromodeoxyuridine (BrdU) proliferation assay. Brachial loading increased DNA synthesis in cells from all three donors, and brachial+E/E inhibitor treatment increased DNA synthesis in two of three donor lines, relative to the static control (
Senescent MSCs have a reduced capacity to differentiate into other cell phenotypes, which limits their clinical utility (Siegel et al., 2013). To test if mechanical conditioning improves the multipotency of aged MSCs, brachial loading and E/E inhibitor treatments were repeated on P5 MSCs from an 81-year-old female donor. Following the treatments, the MSCs were expanded in culture until P8, then evaluated for adipogenic and osteogenic differentiative potential with the hMSC functional identification kit (R&D Systems, SC006). A young MSC line from a 24-year-old female donor was also expanded until P8 and included as a control. The rationale for using cells at P8 was that fully senescent MSCs are expected to completely lose the ability to undergo functional tri-linage differentiation, and studies have shown that an almost complete loss in multipotency occurs around P9/P10 for cells derived from old donors (60+ years) (Zaim et al., 2012). Thus, cells were tested just prior to the theoretic point of complete senescence to determine if the mechanical and pharmaceutical conditioning treatments can improve impaired multipotency. Following differentiation, cells were fixed and stained with FABP4 to evaluate adipogenic differentiation and Osteocalcin to evaluate osteogenic differentiation (
The Sirtuin family of proteins are nicotinamide dinucleotide (NAD+) dependent enzyme deacylases that regulate many cellular processes, including DNA repair, differentiation, metabolism, inflammation, and aging (Lee et al., 2019). Several members of the Sirtuin family have also been implicated as major regulators of multiple senescence pathways (Cakouros & Gronthos, 2019; Onn et al., 2020). To investigate the effects of mechanical and pharmaceutical conditioning on Sirtuin expression, MSCs from an 81-year-old donor were treated with brachial loading and E/E inhibition and Western blotting performed for several proteins in the Sirtuin family. Results showed significant upregulation of SIRT1, p-SIRT1, a SIRT2 isoform, and SIRT6 with brachial loading, and upregulation of p-SIRT1, SIRT6, and SIRT7 with brachial+E/E treatment, in comparison to the static control (
The effective management of oxidative stress and repair of DNA damage are critically important mechanisms for preventing senescence in MSCs (Banimohamad-Shotorbani et al., 2020; Brandl et al., 2011; Yu et al., 2018). To test if the mechanical and pharmaceutical treatments could alter these mechanisms, the mechanical and pharmaceutical treatments were repeated on MSCs from an aged donor (age 81), which were evaluated for protein expression with Western blotting in pathways related to oxidative stress (
Long term culture of MSCs impairs the ATM-dependent recognition of DNA breaks, leading to the buildup of DNA damage and genetic instability (Hladik et al., 2019). To investigate the effects of mechanical loading on DNA damage repair signaling, the mechanical and pharmaceutical treatments were repeated on MSCs from an aged donor (age 81), which were evaluated for protein expression by Western blotting in pathways related to DNA damage repair (
To investigate the downstream effects of mechanically altered DNA damage repair signaling, MSCs from the 81-year-old donor were treated with brachial loading or static conditions for 30 minutes, and 4 hours. Next, immunostaining for p-γ-H2AX DNA double-strand break repair foci was performed. The histone H2AX makes a critical contribution to genetic stability through signaling the recognition of DNA damage events and acting as a foundation for the assembly of repair machinery (Pinto & Flaus, 2010). Brachial treatment for 30 minutes resulted in a significantly higher count of p-γ-H2AX foci compared to the static control at the same time point. However, after 4 hours of brachial loading, p-γ-H2AX foci were significantly reduced compared to the 4-hour static control group (
The nuclear morphology of the mechanically treated MSCs was analyzed with the Nuclear Irregularity Index (NII) plugin in FIJI (Filippi-Chiela et al., 2012). MSCs from a young donor (24 year old female) or an aged donor (81 year old female) were treated with brachial loading or static control treatment for 4 hours. Cells were immunostained for DAPI and processed with the NII plugin to characterize their nuclei as “Normal”, “Apoptotic”, “Senescent”, or “Irregular” (
To further investigate the mechanism by which mechanical loading enhances the recognition and repair of DNA damage, the mechanical treatment of aged MSCs was repeated in combination with several pharmaceutical inhibitors of nuclear mechanotransductive signaling. Previous studies have demonstrated that the dynamic tensile loading of MSCs results in altered chromatin condensation that persists long after the completion of mechanical conditioning (Heo et al., 2015). This phenomena, deemed “mechanical memory”, may provide rationale for the long-term functional benefits demonstrated in this study after mechanical conditioning of aged MSCs. Previous work identified that mechanically driven changes in MSC chromatin architecture are dependent on Piezo mechanosensitive calcium channel signaling (Heo et al., 2015). To investigate the role of mechanosensitive signaling in enhanced DNA damage repair, MSCs from an aged donor were treated with brachial loading or static control treatment for 4 hours and co-treated with 40 μM Importazole, a drug that blocks importin-β-dependent nuclear import (Owens et al., 2020), 2 μM Dooku 1, which blocks Yoda1-induced activation of the Piezo1 Ca2+ mechanosensitive ion channel (Evans et al., 2018), or GsMTx4, which directly inhibits Piezo1 channel activity (Mousawi et al., 2020). Following the treatments, MSCs were immunostained for p-γ-H2AX foci (
Next, the role of oxidative stress signaling and ATM-mediated DNA damage repair in mechanically rejuvenating the proliferative function of aged MSCs, and protecting them from senescence, was evaluated. MSCs were treated with 5 μM KU55933 (an ATM kinase inhibitor, ATMi) (Wu et al., 2017), 5 mM N-acetyl-L cysteine (NAC), which has been shown to have strong antioxidant effects in MSCs (Watanabe et al., 2018), or control treatments. Cells were then mechanically conditioned and evaluated for DNA synthesis with a BrdU assay. Results showed significantly higher DNA synthesis under brachial loading and brachial+KU55933 treatment, but not under brachial+NAC treatment (
Protein expression in pathways related to oxidative stress and DNA damage repair was also evaluated by Western blotting (
To investigate the effects of mechanical conditioning on gene expression patterns that are correlated with senescence in MSCs, MSCs were obtained from one aged donor (81 years old) and conditioned with brachial loading or static control treatment. Following the treatments, RNA-seq analysis was performed on the cells. A differential gene expression analysis revealed that brachial mechanical loading treatment significantly regulated 183 genes in comparison to the static control treatment (
Next, a gene set enrichment analysis (GSEA) was performed on the RNA-seq data. The enrichment scores were calculated comparing the genes expressed between mechanically conditioned cells and control cells for gene sets upregulated in cell senescence (CellAge) (Avelar et al., 2020), upregulated in upregulated in elderly donor derived MSCs (E-MTAB-4879) (Peffers et al., 2016), or and upregulated in high passage MSCs (GSE137186) (Salerno et al., 2020). In all three senescence or aging-related gene sets, a significant reduction in gene enrichment was observed with brachial mechanical loading (
To investigate the mechanical loading effect on altered chromatin accessibility, MSCs were obtained from one aged donor (81 years old) and conditioned with brachial loading or static control treatment. Following the treatments, an assay for transposase-accessible chromatin using sequencing (ATAC-seq) was performed on the cells. The enrichment of accessible chromatin at genomic features between static and brachial treated cells was compared (
Cell lines and cell culture. Human mesenchymal stem cells (Promocell) were cultured in low glucose DMEM medium supplemented with 15% fetal bovine serum, L-glutamine and penicillin/streptomycin. Following trypsinization, cells were seeded on the membranes at 20,000 cells per cm2 before mechanical loading. The cells were used between passages 4-6, unless otherwise specified. The MSCs were derived from a donor 1 (Caucasian female, 81 years old), unless otherwise specified. A subset of the studies were performed using MSCs from donor 2 (Caucasian male, 63 years old), or donor 3 (Caucasian male, 68 years old), or donor 4 (Caucasian female, 92 years old). A subset of the studies used MSCs derived from a young, healthy donor (Caucasian female, 24 years old). Human umbilical vein endothelial cells (HUVECs; PromoCell GmbH) were cultured in MCDB 131 medium supplemented with 7.5% fetal bovine serum, L-glutamine and penicillin/streptomycin and EGM-2 SingleQuot Kit (Thermo Fisher Scientific, Inc.). All cells were cultured in an incubator at 37° C. under a 5% CO2 atmosphere.
Application of biomechanical treatment. Mechanical strain was applied to cell culture using a high throughput system described previously (Lee et al., 2013; Lee & Baker, 2015; Lee et al., 2020). Briefly, cells are cultured on custom made plates that are mounted on a system that applies strain. The cell culture plates were comprised of silicone membranes (0.005″ thickness; Specialty Manufacturing, Inc.) that are sandwiched between two plates, with silicone rubber gaskets at the interfaces to prevent leaking. These cell culture membranes are UV sterilized and coated with 50 μg/mL fibronectin overnight at 37° C. to allow cell adhesion. After cell attachment, the plates are mounted on to the top plate of the system using screws. To apply mechanical strain, a platen with 36 Teflon pistons is moved into the cell culture membrane. The motion is driven by a hygienically sealed, voice coil-type linear motor (Copley Controls). The platen is stabilized using six motion rails mounted with linear motion bearings. The hygienically sealed motor housing has chilled water running through in order to prevent overheating during operation. Unless otherwise specified, loading was applied for 4 hours per day for 7 days with or without 1 μm the EGFR/ErbB-2/ErbB-4 Inhibitor (CAS 881001-19-0; Table 3).
BrdU Proliferation Assay. Following the mechanical and chemical treatments, cells were removed from the stretch plates using 0.05% Trypsin-EDTA, seeded onto optical-bottom 96 well plates at a density of 10,000 cells/well, and allowed to grow for 24 hours. Bromodeoxyuridine (5-bromo-2′-deoxyuridine; BrdU) labeling solution was added to culture media. Cells were washed and stained with the detection and HRP antibodies, following the BrdU assay protocol according to the manufacturer's instructions (Cell Signaling Technology). Absorbance was read at 450 nm using a FlexStation-3 plate reader (Molecular Devices).
Long-term culture expansion assay. Following the mechanical and chemical treatments, cells were removed from the stretch plates using 0.05% Trypsin-EDTA and counted with a hemacytometer. Cells were seeded into standard 6 well culture plates (Corning) at a density of 100,000 cells/well, with an n=2 for each treatment group. Every 7 days, cells were trypsinized, counted, and reseeded into new 6 well plates at an equal cell seeding density. At the end of the study, cumulative doubling events across all passages were calculated for each treatment group.
Cell lysis and immunoblotting. Following the treatments, the cells were lysed in 20 mM Tris with 150 mM NaCl, 1% Triton X-100, 0.1% SDS, 2 mM sodium orthovandate, 2 mM PMSF, 50 mM NaF and a protease inhibitor cocktail (Roche, Inc.). The proteins were separated on a NuPAGE 10% bis-tris midi gel (Novex) and transferred to nitrocellulose membrane using iBlot transfer stack (Novex). The membranes were blocked for one hour in 5% non-fat milk in PBS with 0.01% tween-20 (PBST). After washing twice in PBST, cells were incubated with primary antibodies (Table 1) overnight in 1% non-fat milk at 4° C. The membranes were washed with PBST and incubated at room temperature for two hours with secondary antibody. The membrane was treated with chemiluminescent substrate (SuperSignal West Femto; Thermo Fisher Scientific, Inc.) then imaged using a digital imaging system (Cell Biosciences, Inc.).
Human mesenchymal stem cell multipotency analysis. MSCs at passage 4 were cultured and treated with biomechanical conditioning as described previously. Following the treatments, MSCs removed from the stretch plates with 0.05% Trypsin-EDTA and seeded onto standard culture plates, then expanded under normal culture conditions until passage 8. Following the protocol from the hMSC functional identification kit (R&D Systems; #SC006), P8 MSCs were removed from the culture plates with 0.05% Trypsin-EDTA and seeded onto glass coverslips in 24 well plates at a density of 2.1×104 cells/cm2 for the adipogenic differentiation and 4.2×103 cells/cm2 for the osteogenic differentiation. Cells were treated with the respectively provided differentiation media every 3 days for a total of 21 days. Following the differentiation, the cells were fixed in 4% paraformaldehyde in PBS for 10 minutes followed by washing with PBS. Next, samples were blocked and permeabilized with PBS containing 5% FBS, 1% BSA, and 0.3% Triton X-100 PBS for 40 minutes. After washing, cells were incubated with primary antibodies (see Table 2 for specific antibodies and concentrations) in PBS with 1% BSA overnight at 4° C. The samples were then washed twice in PBS with 1% BSA and incubated with secondary antibodies in PBS with 1% BSA for 2 hours in a light protected environment. Cells treated with extensive washes with PBS with 1% BSA prior to flipping the coverslip and mounting on glass slides in anti-fade media (Vector Laboratories, Inc.). The samples were then imaged using confocal fluorescence microscopy (LSM 710 Confocal; Carl Zeiss, Inc.).
Reactive oxygen species assay. MSCs were treated with brachial mechanical loading for 2 hours. Next, culture media was removed, and chemical treatments were added to fresh cell media. Cells received hydrogen peroxide (H2O2; 0.4 mM in PBS), N-acetyl-cysteine (NAC; 5 mM in culture media), or fresh culture media only. The mechanical loading was resumed for 1 additional hour, and chemical treatments were replaced with fresh culture media. The mechanical loading was then resumed for an additional 30 minutes. 4 μL of CellRox DeepRed reagent (ThermoFisher Scientific) was added directly to the cell culture media in each well, and the mechanical loading was resumed for an additional 30 minutes. Following the loading, cells were washed with PBS, removed from the stretch plates using 0.05% Trypsin-EDTA, and centrifuged for 3 minutes at 300G, and supernatant removed. Cells from each treatment group were resuspended in 1 mL room temperature stain buffer (BD Biosciences) in flow cytometry tubes (Falcon). Cell suspensions were centrifuged for 3 minutes at 300G, supernatant removed, and resuspended in 1 mL of stain buffer with 1 μL of Sytox Blue Live/Dead reagent (ThermoFisher Scientific) and incubated for 10 minutes at room temperature in a light-protected environment. Cell suspensions were centrifuged, supernatant removed, and resuspended in 1 mL of 1× fix/perm solution (BD Biosciences) and incubated at 4° C. for 30 minutes. Following fixation, cells were centrifuged with washing buffer two more times, before resuspended in stain buffer and measured. A BD LSR II Fortessa Flow Cytometer (BD Biosciences) was used to measure population fluorescent signals. At least 10,000 events were recorded and further gating and quantification was done through FlowJo software.
Immunocytochemical staining on silicone membranes. Following the treatments, the cells were fixed in 4% paraformaldehyde in PBS for 10 minutes followed by washing and permeabilization with 0.1% Triton X-100 PBS for 5 minutes. Next, samples were blocked with PBS containing 5% FBS and 1% BSA for 40 minutes. After washing, cells were incubated with primary antibodies (see Table 2 for specific antibodies and concentrations) in PBS with 1% BSA overnight at 4° C. The samples were then washed twice in PBS with 1% BSA and incubated with secondary antibodies in PBS with 1% BSA for 2 hours in a light protected environment. Cells treated with extensive washes with PBS with 1% BSA prior to mounting in anti-fade media (Vector Laboratories, Inc.). The samples were then imaged using confocal microscopy with a LSM 710 laser scanning confocal microscope (Carl Zeiss, Inc.).
p-γ-H2AX Foci Analysis. Following confocal imaging, images were analyzed with FIJI software to count p-γ-H2AX histone foci. Three-dimensional image stacks were maximum intensity projected to generate two-dimensional images with 12-bit range of 0-4095. Background signal outside of nuclei boundaries was measured and subtracted from the image. Individual cell nuclei regions of interest were selected, and an intensity threshold of 300 bits was applied. The Analyze Particle plugin was applied to count the p-γ-H2AX foci in each nucleus, with a minimum particle size of 0.5 μm applied. At least 100 individual cell nuclei were analyzed for each treatment group to calculate the mean foci count per cell, mean foci count per nuclear area, and standard errors.
Nuclear Morphometric Analysis. Following confocal imaging, DAPI images were analyzed with the Nuclear Irregularity Index (NII) plugin in FIJI to analyze the nuclear morphology of the cells (Filippi-Chiela et al., 2012). Nuclei were manually marked by defining the nuclei intensity segmentation threshold, as described in the NII plugin manual. Images of nuclei from a young, healthy (24-year-old) MSC line were used to parametrize the “normal nuclei”. Images of nuclei from brachial or static control treated MSCs were included as “treated nuclei”, with at least 100 nuclei analyzed for each group. Nuclei were classified as “Normal”, “Apoptotic”, “Senescent”, or “Irregular” based on measurements of their area, area/box, aspect, radius ratio, and roundness, in comparison to the young “normal nuclei” control.
Comet assay. Cells were treated with brachial mechanical loading at 0.1 Hz and 7.5% maximum strain for 30 minutes, 4 hours, cultured under static conditions, or cultured under static conditions with 0.4 mM H2O2 to induce DNA damage. Following the treatments, a subset of the cells were left in static conditions for 20 hours. Next, a neutral comet assay was performed to quantify the DNA double-strand-breaks, using the Comet Assay Kit (Abcam), and following the manufacturer's protocol. Briefly, the lysis buffer, alkaline solution, and TBE neutral electrophoresis solution were prepared in advance and stored at 4° C. Comet agarose solution was heated at 90° C. for 20 minutes, stirred until homogenized, and pipetted onto the comet slides, then stored horizontally at 4° C. Following the treatments, culture media was removed from the cells, then cells were washed with dPBS and incubated with 25 mM EDTA for 10 minutes at 37° C. Cells were gently removed with pipetting, centrifuged, and resuspended at 2×105 cells/mL in 4° C. dPBS. Cell suspensions were combined with molten comet agarose at a 1/10 (v/v) ratio. 75 μL of cell/agarose suspensions was pipetted into each well of the comet slides, on top of the agarose base layer. Slides were allowed to chill at 4° C. until the agarose solidified. Next, slides were incubated with 4° C. lysis buffer for 60 minutes in light-protected conditions. Lysis buffer was aspirated and replaced with 4° C. alkaline solution and incubated for 30 minutes in light-protected conditions. Alkaline solution was aspirated and replaced with 4° C. TBE electrophoresis solution, then slides were carefully transferred to a horizontal electrophoresis chamber (BioRad). 20 volts were applied for 20 minutes, then slides were transferred to 4° C. deionized water for washing. Lastly, slides were fixed with 4° C. 70% ethanol, then air dried. 1× Vista Green DNA Dye (Abcam) was added to the slides and incubated at room temperature for 15 minutes. The samples were then imaged using confocal microscopy with a LSM 710 laser scanning confocal microscope (Carl Zeiss, Inc.). The OpenComet plugin in FIJI was used to score the comets and quantify the Olive Tail Moment (OTM) for each treatment group (Gyori et al., 2014), with at least 100 comets scored per group.
Conditioning of hMSCs using biochemical factors. For mechanistic experiments involving chemical factors and biological inhibitors, cells were incubated with the chemical treatments as shown in Table 3. The cells were treated with mechanical loading for 4 hours/day under brachial waveform at 0.1 Hz and 7.5% maximum strain or cultured under static conditions. The culture media containing the treatments were replaced on day 3 and day 5 for all 7-day treatments.
β-Galactosidase Analysis. Following the treatments, cell culture media was removed, and cells were washed once with PBS. Cells were fixed and stained for β-Galactosidase following the protocol of the Senescence β-Galactosidase Staining Kit (Cell Signaling Technologies). Following staining, cells were stored with 70% glycerol and imaged using a light microscope (Meiji, Inc.). Images were post-processed in Adobe Photoshop to increase image contrast and visibility of β-Galactosidase+ cells. Total cells and β-Galactosidase+ cells were manually counted in FIJI, with at least 1,000 cells counted per treatment group.
Flow Cytometry. For the characterization of mesenchymal stem cells during long-term culture expansion (passage 5-10), the cells were detached from the culture plate using Accutase (Sigma-Aldrich) and were labeled with fluorescent antibodies according to the R&D Systems human mesenchymal stem cell verification flow kit protocol (R&D FMC020; see Table 4 for the specific antibodies used). Briefly, the detached cells were centrifuged and the supernatant was removed. Fixing and permeabilizing buffer was added while the cells were vortexed and incubated for 40 min. Next, the samples were centrifuged and the supernatant was removed. Cells were then treated with washing buffer containing antibodies for 50 min. Following antibody incubation, cells were centrifuged with washing buffer two more times, then were treated with stain buffer and measured. A BD LSR II Fortessa Flow Cytometer (BD Biosciences) was used to measure population fluorescent signals. At least 10,000 events were recorded and further gating and quantification was done through FlowJo software.
RNA Sequencing and Analysis. Following treatments, RNA was isolated from the cells from four independent wells per group using the Qiagen RNeasy Mini Kit. RNAseq was performed using an Illumina HiSeq 2500 sequencing machine. For sequencing, single reads of 50 base pairs were performed after poly-A mRNA capture using the Poly(A) Tailing Kit (Ambion) and Ultra II Directional RNA Library Prep Kit (NEB) to isolate mRNA and perform dUTP directional preparation of the mRNA library. RNA sequencing was performed by GENEWIZ (Azenta Life Sciences). Gene expression analysis was performed using R. Gene ontology, Pathview, and Kyoto Encyclopedia of Genes and Genomes (KEGG) analysis was performed using the Molecular Signatures Database (En), PANTHER, or ShinyGO (Kanehisa et al., 2021; Luo & Brouwer, 2013; Ge et al., 2020).
Gene set enrichment analysis (GSEA). To define signature gene sets of senescent cells, aged-donor-derived MSCs, or high passage MSCs, we downloaded multiple RNA-seq data sets (Table 5). Obtained RNA-seq data sets were mapped to the human transcripts (GRCh38) with Salmon mapper (v1.1.0) (Patro et al., 2017) and the count of mapped reads was normalized as transcripts per million (TPM) using an R package of tximport (v 1.14.2) (Soneson et al., 2015). Multiple processed RNA-seq data belonging to each cell-type category were considered as replicates and compared to data obtained from mesenchymal stem cells (MSCs) using edgeR (v 3.26.8) (Robinson et al., 2010) to define signature genes for each cell-type. GSEA was performed using
Statistical analysis. All results are shown as mean±standard error of the mean. All experiments used biological replicates that consisted of cells in non-repeated, independent cell culture wells or tissue samples from different animals, unless specified otherwise. Multiple comparisons between groups were analyzed by two-way ANOVA followed by a Tukey post-hoc or a Dunnett post-hoc test when testing multiple comparisons versus a control group. For non-parametric data, multiple comparisons were made using the Kruskal-Wallis test followed by post-hoc testing with the Conover-Iman procedure. A p-value of 0.05 or less was considered statistically significant for all tests.
All of the methods disclosed and claimed herein can be made and executed without undue experimentation in light of the present disclosure. While the compositions and methods of this invention have been described in terms of preferred embodiments, it will be apparent to those of skill in the art that variations may be applied to the methods and in the steps or in the sequence of steps of the method described herein without departing from the concept, spirit and scope of the invention. More specifically, it will be apparent that certain agents which are both chemically and physiologically related may be substituted for the agents described herein while the same or similar results would be achieved. All such similar substitutes and modifications apparent to those skilled in the art are deemed to be within the spirit, scope and concept of the invention as defined by the appended claims.
The following references, to the extent that they provide exemplary procedural or other details supplementary to those set forth herein, are specifically incorporated herein by reference.
The present application claims the priority benefit of U.S. provisional application No. 63/582,416, filed Sep. 13, 2023, the entire contents of which are incorporated herein by reference.
This invention was made with government support under Grant no. R01 HL141761 awarded by the National Institutes of Health. The government has certain rights in the invention.
| Number | Date | Country | |
|---|---|---|---|
| 63582416 | Sep 2023 | US |
