Patent Application
20220249627
Incorporated herein by reference in its entirety is the sequence listing submitted via EFS-Web as a text file named SEQLIST.txt, created Feb. 15, 2022, and having a size of 135,051 bytes.
The present invention relates to the fields of biofilm deposition and the treatment of disease. More specifically, the invention provides compositions and methods useful for the treatment of dental caries and other oral diseases. The invention also provides methods for coating biomedical devices for inhibiting undesirable biofilm deposition thereon.
Several publications and patent documents are cited throughout the specification in order to describe the state of the art to which this invention pertains. Each of these citations is incorporated by reference herein as though set forth in full.
Biopharmaceuticals produced in current systems are prohibitively expensive and are not affordable for large majority of the global population. The cost of protein drugs ($140 billion in 2013) exceeds GDP of >75% of countries around the globe [Walsh 2014], making them unaffordable. One third of the global population earns <$2 per day and can't afford any protein drug (including the underprivileged, elderly and lower socio-economic groups in the US). Such high costs are associated with protein production in prohibitively expensive fermenters, purification, cold transportation/storage, short shelf life and sterile delivery methods [Daniell et al 2015, 2016].
Biofilms are formed by a complex group of microbial cells that adhere to the exopolysaccharide matrix present on the surface of medical devices. Biofilm-associated infections associated with medical device implantation pose a serious problem and adversely affects the function of the device. Medical implants used in oral and orthopedic surgery are fabricated using alloys such as stainless steel and titanium. Surface treatment of medical implants by various physical and chemical techniques has been attempted in order to improve surface properties, facilitate biointegration and inhibit bacterial adhesion as bacterial adhesion is associated with surrounding tissue damage and often results in malfunction of the implant.
Many infectious diseases in humans are caused by biofilms, including those occurring in the mouth [Hall-Stoodley et al., 2004; Marsh, et al 2011]. For example, dental caries (or tooth decay) continues to be the single most prevalent biofilm-associated oral disease, afflicting mostly underprivileged children and adults in the US and worldwide, resulting in expenditures of >$81 billion annually [Beiker and Flemmig, 2011; Dye et al., 2015; Kassebaum et al, 2015]. Caries-causing (cariogenic) biofilms develop when bacteria accumulate on tooth-surfaces, forming organized clusters of bacterial cells that are firmly adherent and enmeshed in an extracellular matrix composed of polymeric substances such as exopolysaccharides (EPS) [Bowen and Koo, 2011].
Additionally, aerosolized microbes generated during dental procedures and mechanical plaque/biofilm removal have been recognized as potential source for the spread of several infectious diseases (Bennett et al., 2000). This has received greater attention during the current COVID-19 global pandemic (Xu et al., 2020). The saliva and dorsum of the tongue are major sources of SARS-CoV-2 and stability of the virus in the aerosol and its spread in the aerosolized form has been widely reported (Van Doremalen et al., 2020; World Health Organization [WHO], 2020; Xu et al., 2020). Therefore, WHO and dental associations including American Dental Association recommended suspension of aerosol-generating procedures in the clinic except for emergencies (Bennett et al., 2000; Van Doremalen et al., 2020; WHO, 2020; Xu et al., 2020).
Furthermore, COVID-19 patients have shown high accumulation of pathogenic oral bacteria, whereas poor oral hygiene which disproportionately afflicts impoverished populations, may be a risk factor for COVID-19 (Patel and Sampson, 2020). Accumulation of microbes on teeth leads to the formation of intractable dental biofilms (plaque) that cause oral diseases such as dental caries (tooth decay) requiring costly clinical interventions at the dental office. Hence, development of alternative methods for plaque control at home is of paramount importance and urgency.
Current topical antimicrobial modalities for controlling cariogenic biofilms are limited. Chlorhexidine (CHX) is considered the ‘gold standard’ for oral antimicrobial therapy, but has adverse side effects including tooth staining and calculus formation, and is not recommended for daily therapeutic use [Jones, 1997; Autio-Gold, 2008]. As an alternative, several antimicrobial peptides (AMPs) have emerged with potential antibiofilm effects against caries-causing oral pathogens such as Streptococcus mutans [da Silva et al., 2012; Guo et al., 2015].
Antimicrobial peptides (AMP) are an evolutionarily conserved component of the innate immune response and are naturally found in different organisms, including humans. When compared with conventional antibiotics, development of resistance is less likely with AMPs. They are potently active against bacteria, fungi and viruses and can be tailored to target specific pathogens by fusion with their surface antigens (Lee et al 2011; DeGray et al 2001; Gupta et al 2015). Linear AMPs have poor stability or antimicrobial activity when compared to AMPs with complex secondary structures. For example, retrocyclin or protegrin have high antimicrobial activity or stability when cyclized (Wang et al 2003) or when forming a hairpin structure (Chen et al 2000) via disulfide bond formation. RC 101 is highly stable at pH 3, 4, 7 and at temperatures ranging from 25° C. to 37° C. as well as in human vaginal fluid for 48 hours (Sassi et al 2011a), while its antimicrobial activity was maintained for up to six months (Sassi et al 2011b). Likewise, protegrin is highly stable in salt or human fluids (Lai et al 2002; Ma et al 2015) but lost potency when linearized. These intriguing characteristics of antimicrobial peptides with complex secondary structures may facilitate development of novel therapeutics. However, the high cost of producing sufficient amounts of antimicrobial peptides as well as other biofilm degrading enzymes is a major barrier for their clinical development and commercialization.
In accordance with the present invention, a multi-component composition comprising at least one antimicrobial peptide (AMP) and at least one biofilm degrading enzyme which act synergistically to degrade biofilm structures and inhibit biofilm deposition is provided. In certain embodiments, the AMP is selected from protegrin 1, RC-101 and the AMPs listed in Table 1. The biofilm degrading enzyme, includes, for example, mutanase, dextranase, glucoamylase, deoxyribonuclease I, DNAase, dispersin B, glycoside hydrolases and the enzymes provided in Table 2. In certain embodiments, the coding sequences for these enzymes are codon optimized for expression in a plant chloroplast. In a particularly preferred embodiment, the at least one AMP and at least one biofilm degrading enzyme are produced recombinantly. In a particularly preferred embodiment, the AMP and biofilm degrading enzyme(s) are expressed as a fusion protein. When the composition is for the treatment of oral diseases, the composition may optionally further comprise an antibiotic, fluoride, CHX or all of the above. The composition may be contained within chewing gum, hard candy, or within an oral rinse. Preferred fusion proteins of the invention include, without limitation, PG-1-Mut, PG-1-Dex, PG-1-Mut-Dex, RC-101-Mut, RC-101-Dex, RC-101-Mut-Dex for use alone or in combination for the degradation of biofilms. Notably any of the AMPs listed in Table 1 can replace either PG-1 or RC-101 in the aforementioned fusion proteins to alter or improve the bacteriocidal action of the fusion protein.
To alter the degradation activity of the fusion proteins, the enzymes listed above and hereinbelow may replace Mut, Dex or both in the fusion proteins of the invention. In another embodiment, when two different EPS enzymes are employed in the compositions, such enzymes may be delivered at different ratios, e.g., 1:2, 1:3, 1:4, 1:5, 1:6, 1:7, 1:8 etc. When Mut and Dex are delivered together in a gum or oral rinse for example, a preferred ratio is 5:1 Dex:Mut.
In another aspect, the invention comprises a biofilm degrading composition harboring a mutanase enzyme. in a pharmaceutically acceptable carrier. In preferred embodiments, the mutanase is encoded by SEQ ID NO: 1 and is expressed in a plant plastid. In certain embodiments, the composition further comprises a plant remnant. The plant remnant may be freeze dried. In a particularly preferred embodiment, the plant remnant is from tobacco or lettuce. In certain embodiments, the composition comprises at least one AMP, lipase, and/or biofilm degrading enzyme. In certain embodiments, the lipase is obtained from a commercial vendor or produced in a plant plastid present in said plant remnant. The biofilm degrading enzyme, includes, for example, dextranase, glucoamylase, deoxyribonuclease I, DNAase, dispersin B, glycoside hydrolases and the enzymes provided in Table 2. The at least one biofilm degrading enzyme may have a sequence selected from SEQ ID NOS: 2, 12, 14, 16, 18, 20, 24, and 26. In particularly preferred embodiments, the composition comprises mutanase, dextranase and lipase present in a chewing gum carrier. In certain embodiments, the dextranase and mutanase are present in a 5:1 ratio in said chewing gum.
In certain embodiments, the biofilm degrading composition further comprises an antimicrobial, an antibiotic, fluoride, and/or CHX. When the composition is for the treatment of oral diseases, the composition may be contained within chewing gum, hard candy, or within an an oral rinse.
In another aspect, the invention provides a method of degrading and/or removing biofilm comprising contacting a surface harboring said biofilm with the compositions described above, the composition having a bactericidal effect, and reducing or eliminating said biofilm comprising one or more undesirable microorganisms, wherein when said biofilm is present in or on an animal subject in need of said reduction or elimination. In certain embodiments, the biofilm is present in the mouth. In other embodiments, the biofilm is present on an implanted medical device. The method may also be used to remove biofilms present in an internal or external body surface is selected from the group consisting of a surface in a urinary tract, a middle ear, a prostate, vascular intima, heart valves, skin, scalp, nails, teeth and an interior of a wound. In particularly preferred embodiments, the composition used in these methods comprises mutanase, dextranase, and lipase in a suitable carrier.
In yet another embodiment, the composition of the invention comprises said at least one AMP and said at least one biofilm degrading enzyme are produced in a plant plastid. The plant may be a tobacco plant and the sequences encoding said AMP and enzyme is codon optimized for expression in a plant plastid. In a preferred embodiment, the AMP and biofilm degrading enzyme are expressed in a lettuce plant as a fusion protein under the control of endogenous regulatory elements present in lettuce plastids. In other embodiments, the composition does not comprise an AMP, but does contain lipase.
Many infectious diseases in humans are caused by virulent biofilms, including those occurring within the mouth (e.g. dental caries and periodontal diseases). Dental caries (or tooth decay) continues to be the single most costly and prevalent biofilm-associated oral disease in the US and worldwide. It afflicts children and adults alike, and is a major reason for emergency room visits leading to absenteeism from work and school. Unfortunately, the prevalence of dental caries is still high (>90% of US adult population) and it remains the most common chronic disease afflicting children and adolescents, particularly from a poor socio-economic background. Furthermore, poor oral health often leads to systemic consequences and impacts overall health. Importantly, the cost to treat the ravages of this disease (e.g. carious lesions and pulpal infection) exceeds $40 billion/yr in the US alone. Fluoride is the mainstay of dental caries prevention. However, its widespread use offers incomplete protection against the disease.
Fluoride is effective in reducing demineralization and enhancing demineralization of early carious lesions, but has limited effects against biofilms. Conversely, current antimicrobial modalities for controlling caries-causing biofilms are largely ineffective.
There is an urgent need to develop efficacious therapies to control virulent oral biofilms. In accordance with the present invention, methods for low-cost production and delivery of therapeutically effective plant-expressed biopharmaceuticals superior to current antibiofilm/anti-caries modalities are provided.
As used herein, antimicrobial peptides are small peptides having any bacterial activity. “RC-101” is an analogue of retrocyclin, a cyclic octadecapeptide, which can protect human CD4+ cells from infection by T- and M-tropic strains of HIV-1 in vitro and prevent HIV-1 infection in human cervicovaginal tissue. The ability of RC-101 to prevent HIV-1 infection and retain full activity in the presence of vaginal fluid makes it a good candidate for other topical microbicide applications, especially in oral biofilms. The sequence of RC-101 is provided in Plant Biotechnol J. 2011 January; 9(1): 100-115 which is incorporated herein by reference.
“C16G2” is a novel synthetic antimicrobial peptide with specificity for S. mutans,
“Protegrin-1 (PG)” is a cysteine-rich, 18-residue β-sheet peptide. It has potent antimicrobial activity against a broad range of microorganisms, including bacteria, fungus, virus, and especially some clinically relevant, antibiotic-resistant bacteria. For example, bacterial pathogens E. coli and fungal opportunist C. albicans are effectively killed by PG in laboratory testing. The sequence of PG-1 is provided in Plant Biotechnol J. 2011 January; 9(1): 100-115 which is incorporated herein by reference.
Additional antimicrobial peptides include those set forth below in Table 1 below.
a Linker regions between targeting and killing peptides are underlined,
b Peptide C-terminal arriidation,
A “biofilm” is a complex structure adhering to surfaces that are regularly in contact with water, consisting of colonies of bacteria and usually other microorganisms such as yeasts, fungi, and protozoa that secrete a mucilaginous protective coating in which they are encased. Biofilms can form on solid or liquid surfaces as well as on soft tissue in living organisms, and are typically resistant to conventional methods of disinfection. Dental plaque, the slimy coating that fouls pipes and tanks, and algal mats on bodies of water are examples of biofilms. Biofilms are generally pathogenic in the body, causing such diseases as dental caries, cystic fibrosis and otitis media.
“Biofilm degrading enzymes” include, without limitation, exo-polysaccharide degrading enzymes such as dextranase, mutanase, DNAse, endonuclease, deoxyribonuclease I, dispersin B, and glycoside hydrolases, such as 1→3)—α-D-glucan hydrolase, although use of chloroplast codon optimized sequences encoding dextranase and mutanase are preferred, the skilled person is well aware of many different biofilm degrading enzymes in the art. Additional enzyme sequences for use in the fusion proteins of the invention are provided below.
As used herein, the terms “administering” or “administration” of an agent, drug, or peptide to a subject includes any route of introducing or delivering to a subject a compound to perform its intended function. The administering or administration can be carried out by any suitable route, including orally, topically, intranasally, parenterally (intravenously, intramuscularly, intraperitoneally, or subcutaneously), or rectally. Administering or administration includes self-administration and the administration by another.
As used herein, the terms “disease,” “disorder,” or “complication” refers to any deviation from a normal state in a subject.
As used herein, by the term “effective amount” “amount effective,” or the like, it is meant an amount effective at dosages and for periods of time necessary to achieve the desired result.
As used herein, the term “inhibiting” or “preventing” means causing the clinical symptoms of the disease state not to worsen or develop, e.g., inhibiting the onset of disease, in a subject that may be exposed to or predisposed to the disease state, but does not yet experience or display symptoms of the disease state.
As used herein, the term “expression” in the context of a gene or polynucleotide involves the transcription of the gene or polynucleotide into RNA. The term can also, but not necessarily, involves the subsequent translation of the RNA into polypeptide chains and their assembly into proteins.
A plant remnant may include one or more molecules (such as, but not limited to, proteins and fragments thereof, minerals, nucleotides and fragments thereof, plant structural components, etc.) derived from the plant in which the protein of interest was expressed. Accordingly, a composition pertaining to whole plant material (e.g., whole or portions of plant leafs, stems, fruit, etc.) or crude plant extract would certainly contain a high concentration of plant remnants, as well as a composition comprising purified protein of interest that has one or more detectable plant remnants. In a specific embodiment, the plant remnant is rubisco.
In another embodiment, the invention pertains to an administrable composition for treating or preventing biofilm formation in situ (e.g., in the mouth) and on biomedical devices useful for surgical implantation such as stents, artificial joints, and the like. In this embodiment, the devices are coated with the composition to inhibit unwanted biofilm deposition on the device. The composition comprises a therapeutically-effective amount of one or more antimicrobial peptides (AMP) and one or more enzymes having biofilm degrading activity in combination, each of said AMP and enzyme thereof having been expressed by a plant and a plant remnant and acting synergisticall to degrade said biofilm. In certain embodiments the AMP(s) and enzymes(s) are expressed from separate plastid transformation vectors. In other embodiments, the plastid transformation vectors comprising polycistronic coding sequences where both the AMP and the enzymes are expressed from a single vector.
Proteins expressed in accord with certain embodiments taught herein may be used in vivo by administration to a subject, human or animal in a variety of ways. The pharmaceutical compositions may be administered orally, topically, subcutaneously, intramuscularly or intravenously, though oral topical administration is preferred.
Oral compositions produced by embodiments of the present invention can be administered by the consumption of the foodstuff that has been manufactured with the transgenic plant producing the plastid derived therapeutic protein. The edible part of the plant, or portion thereof, is used as a dietary component. The therapeutic compositions can be formulated in a classical manner using solid or liquid vehicles, diluents and additives appropriate to the desired mode of administration. Orally, the composition can be administered in the form of tablets, capsules, granules, powders, gums, and the like with at least one vehicle, e.g., starch, calcium carbonate, sucrose, lactose, gelatin, etc. The therapeutic protein(s) of interest may optionally be purified from a plant homogenate. The preparation may also be emulsified. The active ingredient is often mixed with excipients which are pharmaceutically acceptable and compatible with the active ingredient. Suitable excipients are, e.g., water, saline, dextrose, glycerol, ethanol or the like and combination thereof. In addition, if desired, the compositions may contain minor amounts of auxiliary substances such as wetting or emulsifying agents, pH buffering agents, or adjuvants. In a preferred embodiment the edible plant, juice, grain, leaves, tubers, stems, seeds, roots or other plant parts of the pharmaceutical producing transgenic plant is ingested by a human or an animal thus providing a very inexpensive means of treatment of disease.
In a specific embodiment, plant material (e.g. lettuce material) comprising chloroplasts expressing AMPs and biofilm degrading enzymes and combinations thereof, is homogenized and encapsulated. In one specific embodiment, an extract of the lettuce material is encapsulated. In an alternative embodiment, the lettuce material is powderized before encapsulation. As mentioned previously, the biofilm degrading proteins may also be purified from the plant following expression.
In alternative embodiments, the compositions may be provided with the juice of the transgenic plants for the convenience of administration. For said purpose, the plants to be transformed are preferably selected from the edible plants consisting of tomato, carrot and apple, among others, which are consumed usually in the form of juice.
According to another embodiment, the subject invention pertains to a transformed chloroplast genome that has been transformed with a vector comprising a heterologous gene that expresses a combination of peptides as disclosed herein.
Of particular interest is a transformed chloroplast genome transformed with a vector comprising a heterologous gene that expresses one or more AMP and biofilm degrading enzyme or a combination thereof, polypeptide. In a related embodiment, the subject invention pertains to a plant comprising at least one cell transformed to express a peptide as disclosed herein.
Reference to genetic sequences herein refers to single- or double-stranded nucleic acid sequences and comprises a coding sequence or the complement of a coding sequence for polypeptide of interest. Degenerate nucleic acid sequences encoding polypeptides, as well as homologous nucleotide sequences which are at least about 50, 55, 60, 65, 60, preferably about 75, 90, 96, or 98% identical to the cDNA may be used in accordance with the teachings herein polynucleotides. Percent sequence identity between the sequences of two polynucleotides is determined using computer programs such as ALIGN which employ the FASTA algorithm, using an affine gap search with a gap open penalty of −12 and a gap extension penalty of −2. Complementary DNA (cDNA) molecules, species homologs, and variants of nucleic acid sequences which encode biologically active polypeptides also are useful polynucleotides.
Variants and homologs of the nucleic acid sequences described above also are useful nucleic acid sequences. Typically, homologous polynucleotide sequences can be identified by hybridization of candidate polynucleotides to known polynucleotides under stringent conditions, as is known in the art. For example, using the following wash conditions: 2×SSC (0.3 M NaCl, 0.03 M sodium citrate, pH 7.0), 0.1% SDS, room temperature twice, 30 minutes each; then 2×SSC, 0.1% SDS, 50° C. once, 30 minutes; then 2×SSC, room temperature twice, 10 minutes each, homologous sequences can be identified which contain at most about 25-30% basepair mismatches. More preferably, homologous nucleic acid strands contain 15-25% basepair mismatches, even more preferably 5-15% basepair mismatches.
Species homologs of polynucleotides referred to herein also can be identified by making suitable probes or primers and screening cDNA expression libraries. It is well known that the Tm of a double-stranded DNA decreases by 1-1.5° C. with every 1% decrease in homology (Bonner et al., J. Mol. Biol. 81, 123 (1973). Nucleotide sequences which hybridize to polynucleotides of interest, or their complements following stringent hybridization and/or wash conditions also are also useful polynucleotides. Stringent wash conditions are well known and understood in the art and are disclosed, for example, in Sambrook et al., MOLECULAR CLONING: A LABORATORY MANUAL, 2nd ed., 1989, at pages 9.50-9.51.
The following materials and methods are provided to facilitate the practice of the present invention.
Streptococcus mutans UA159 serotype c (ATCC 700610), Actinomyces naeslundii ATCC 12104, Streptococcus gordonii DL-1 and Candida albicans SC5314 were used in present study. These strains were selected because S. mutans is a well-established, virulent cariogenic bacteria [Ajdić D et al, 2002]. S. gordonii is a pioneer colonizer of dental biofilm, and A. naeslundii is also detected during the early stages of dental biofilm formation and may be associated with development of root caries [Dige I et al, 2009]. C. albicans is a fungal organism that colonizes human mucosal surfaces, and it is also detected in dental plaque from toddlers with early childhood caries [Hajeshengallis E et al, 2015]. All strains were stored at −80° C. in tryptic soy broth containing 20% glycerol. Blood agar plates were used for cultivating S. mutans, S. gordonii and A. naeslundii. Sabouraud agar plates were used for C. albicans. All these strains were grown in ultra-filtered (10 kDa molecular-weight cut-off membrane; Prep/Scale, Millipore, MA) buffered tryptone-yeast extract broth (UFTYE; 2.5% tryptone and 1.5% yeast extract, pH 7.0) with 1% glucose to mid-exponential phase (37° C., 5% CO2) prior to use.
The transplastomic plants expressing GFP fused with CTB, PTD, retrocyclin and protegrin were created as described in previous studies [Limaye et al 2006; Kwon et al 2013; Xiao et al 2016; Lee et al 2011]. Transplastomic lines expressing GFP fusion proteins were confirmed using Southern blot assay as described previously [Verma et al 2008]. Also, expression of GFP tagged proteins were confirmed by visualizing green fluorescence from the leaves of each construct under UV illumination.
Purification of GFP fusions Protegrin-1 (PG1) and Retrocyclin (RC101) from transplastomic tobacco was accomplished by organic extraction followed by hydrophobic chromatography done previously (Lee et al, 2011). About 0.2-1 gm of lyophilized leaf material was taken and reconstituted in 10-20 ml of plant extraction buffer (0.2M Tris HCl pH 8.0, 0.1M NaCl, 10 mM EDTA, 0.4M sucrose, 0.2% Triton X supplemented with 2% Phenylmethylsulfonylfluoride and 1 protease inhibitor cocktail). The resuspension was incubated in ice for 1 hour with vortex homogenization every 15 min. The homogenate was then spun down at 75000 g at 4° C. for 1 hour (Beckman LE-80K optima ultracentrifuge) to obtain the clarified lysate. The lysate was subjected to pretreatment with 70% saturated ammonium sulfate and ¼th volume of 100% ethanol, followed by vigorous shaking for 2 min (Yakhnin et al, 1998). The treated solution was spun down at 2100 g for 3 min. The upper ethanol phase was collected, and the process was repeated with 1/16th volume of 100% ethanol. The pooled ethanol phases were further treated with ⅓rd volume of 5M NaCl and ¼th volume of 1-butanol, homogenized vigorously for 2 min and spun down at 2100 g for 3 min. The lowermost phase was collected and loaded onto a 7 kDa MWCO zeba spin desalting column (Thermo scientific) and desalted as per manufacturer's recommendations.
The desalted extract was then subjected to hydrophobic interaction chromatography during the capture phase for further purification. The desalted extract was injected into a Toyopearl butyl—650S hydrophobic interaction column (Tosoh bioscience) which was run on a FPLC unit (Pharmacia LKB-FPLC system). The column was equilibriated with 2.3 column volumes of salted buffer (10 mM Tris-HCl, 10 mM EDTA and 50% saturated ammonium sulfate) to a final 20% salt saturation to facilitate binding of GFP onto the resin. This was followed by a column wash with 5.8 column volumes of salted and unsalted buffer mix and then eluted with unsalted buffer (10 mM Tris-HCl, 10 mM EDTA). The GFP fraction was identified based on the peaks observed in the chromatogram and collected. The collected fractions were subjected to a final polishing step by overnight dialysis. After dialysis the purified proteins were lyophilized (labconco lyophilizer) in order to concentrate the finished product and then stored in −20° C.
Quantification of GFP-RC101 and GFP-PG1 was done by both western blot and fluorescence-based methods. The lyophilized purified proteins were resuspended in sterile 1X PBS and the total protein was determined by Bradford method. The purified protein was then quantified by SDS-PAGE method by loading denatured protein samples along with commercial GFP standards (Vector labs) onto a 12% SDS gel and then western blotting was done using 1:3000 dilution of mouse Anti-GFP antibody (Millipore) followed by probing with 1:4000 dilution of secondary HRP conjugated Goat-Anti Mouse antibody (Southern biotech).
The purified proteins were also quantified using GFP fluorescence. The protein samples were run on a 12% SDS gel under native conditions. After the run, the gel was placed under a UV lamp and then photographed. The GFP concentration in both western and native fluorescence methods was determined by densitometric analysis using Image J software with commercial GFP standards in order to obtain the standard curve. Purity was determined based on GFP quantitation with respect to total protein values determined in Bradford method.
As previously described (Xiao, et al 2016), to determine the uptake of four tags, CTB, PTD, PG1 and RC101, in different human periodontal cell lines, including human periodontal ligament stem cells (HPDLS), maxilla mesenchymal stem cells (MMS), human head and neck squamous cell carcinoma cells (SCC-1), gingiva-derived mesenchymal stromal cells (GMSC), adult gingival keratinocytes (AGK) and osteoblast cells (OBC), briefly, each human cell line cells (2×104) were cultured in 8 well chamber slides (Nunc) at 37° C. overnight, followed by incubation with purified GFP-fused tags: CTB-GFP (8.8 μg), PTD-GFP (13 μg), GFP-PG1 (1.2 μg) and GFP-RC101 (17.3 μg) in 100 μl PBS supplemented 1% FBS at 37° C. for 1 hour. After fixing with 2% paraformaldehyde at RT for 10 min and washing with PBS for three times, all cells were stained with antifade mounting medium with DAPI (Vector laboratories, Inc). For negative control, cells were incubated with commercial GFP (2 μg) in PBS with 1% FBS at 37° C. for 1 hour. All fixed cells were imaged using confocal microscopy. The images were observed under 100× objective, and at least 10-15 GFP-positive cells were recorded for each cell line in three independent analyses.
The killing kinetics of AMPs (Gfp-PG1 and Gfp-RC101) against S. mutans were analyzed by time-lapse killing assay. S. mutans were grown to log phase and diluted to 105 CFU/ml in growth medium. GFP-PG1 and GFP-RC101 were added to S. mutans suspensions at concentrations of 0 to 10 μg/ml and 0 to 80 μg/ml, respectively. At 0, 1, 2, 4, 8 and 24 h, samples were taken and serially diluted in 0.89% NaCl, then spread on agar plates and colonies were counted after 48 h. Absorbance at 600 nm was also checked at each time point. S. gordonii, A. naeslundii and C. albicans suspensions were mixed with Gfp-PG1 at concentration of 10 μg/ml, and at 0, 1 and 2 h, aliquots were taken out for enumeration of CFU.
The effects of AMP on the viability of S. mutans cells were also assessed by time-lapsed measurements. S. mutans were grown to log phase and harvested by centrifugation (5500 g, 10 min) and the pellet was washed once with sodium phosphate-buffered saline (PBS) (pH 7.2), re-suspended in PBS and adjusted to a final concentration of 1×105 CFU/ml. GFP-PG1 was added to S. mutans suspensions at concentrations of 10 μg/ml and 2.5 μM propidium iodide-PI (Molecular Probe Inc., Eugene, Oreg., USA) was added for labeling dead cells. 5 μl of mixtures were loaded on an agarose pad for confocal imaging. Confocal images were acquired using Leica SP5-FLIM inverted single photon laser scanning microscope with a 100X (numerical aperture, 1.4) Oil immersion objective. The excitation wavelengths were 488 nm and 543 nm for GFP and PI, respectively. The emission filter for GFP was a 495/540 OlyMPFC1 filter, while PI was a 598/628 OlyMPFC2 filter. For the time-lapse series, images in the same field of view were taken at 0, 10, 30, and 60 min and created by ImageJ 1.44 (on the world wide web at rsbweb.nih.gov/ij/download.html).
Morphological observations of S. mutans treated with AMP were also examined by scanning electron microcopy (SEM). S. mutans were grown to log phase and diluted to 105 CFU/ml in PBS. Bacteria suspension was mixed with GFP-PG1 (final concentration of 10 μg/ml) for 1 h at 37° C. After treatment, the bacteria were collected by filtration using 0.4 μm Millipore filters. The deposits were fixed in 2.5% glutaraldehyde and 2.0% paraformaldehyde in 0.1 M cacodylate buffer (pH 7.4) for 1 hour at room temperature and processed for SEM (Quanta FEG 250, FEI, Hillsboro, Oreg.) observation. Untreated or bacteria treated with buffer only served as controls.
A well-characterized EPS-matrix producing oral pathogen, S. mutans UA159, and an opportunistic fungal pathogen, C. albicans SC5314 were used to form single- or mixed-speces biofilms on saliva-coated hydroxyapatite disc surfaces. Briefly, hydroxyapatite discs (1.25 cm in diameter, surface area of 2.7±0.2 cm2, Clarkson, Chromatography Products, Inc., South Williamsport, Pa.) were coated with filter-sterilized, clarified human whole saliva (sHA) [Xiao J et alo, 2012]. S. mutans was grown in UFTYE medium with 1% (w/v) glucose to mid-exponential phase (37° C., 5% CO2). Each sHA disc was inoculated with 105 CFU of actively growing S. mutans cells per ml in UFTYE medium containing 1% (w/v) sucrose, and inoculated at 37° C. and 5% CO2 for 19 h. Before inoculum, the sHA discs were topically treated with GFP-PG1 solution (10 ug) for 30 min. The inhibition effect of GFP-PG1 treatment on 3D biofilm architectures were observed via confocal imaging. Briefly, EPS was labeled using 2.5 μM Alexa Fluor 647-labeled dextran conjugate (10 kDa; 647/668 nm; Molecular Probes Inc.), while the bacteria and fungal cells were stained with 2.5 μM SYTO9 (485/498 nm; Molecular Probes Inc.) and Concanavalin A-tetramethyl rhodamine conjugate (Molecular Probes). The imaging was performed using Leica SP5 microscope with 20X (numerical aperture, 1.00) water immersion objective. The excitation wavelength was 780 nm, and the emission wavelength filter for SYTO 9 was a 495/540 OlyMPFEC1 filter, while the filter for Alexa Fluor 647 was a HQ655/40M-2P filter. The confocal image series were generated by optical sectioning at each selected positions and the step size of z-series scanning was 2 μm. Amira 5.4.1 software (Visage Imaging, San Diego, Calif., USA) was used to create 3D renderings of biofilm architecture [Xiao J et al. 2012, Koo H et al. 2010].
To examine the effects of the PG1 on biofilms formed with S. mutans for 19 h on sHA discs, we examined the 3D architecture of the EPS-matrix and in situ cell viability using time-lapse confocal microscopy following biofilms incubation with 1) Control, 2) EPS-degrading enzymes only, 3) PG1 only, or 4) PG1 and EPS-degrading enzymes for up to 60 minutes. The EPS-degrading enzymes used here were dextranase and mutanase, which were capable of digesting the EPS derived from S. mutans by hydrolyzing α-1,6 glucosidic linkages and α-1,3 glucosidic linkages [Hayacibara et al. 2004]. Dextranase produced from Penicillium sp. was commercially purchased from Sigma (St. Louis, Mo.) and mutanase produced from Trichoderma harzianum was kindly provided by Dr. William H. Bowen (Center for Oral Biology, University of Rochester Medical Center). Dextranase and mutanase were mixed at ratio of 5:1 before applying to biofilms [Mitsue F. Hayacibara et al. 2004]. In addition, lipase, dextranase and mutanase were also tested using similar topical regimen as depicted in
At selected time point (19 h), biofilms were removed, homogenized via sonication and subject to microbiological analyses as detailed previously [Xiao J et al. 2012, Koo H et al. 2010]; our sonication procedure does not kill bacteria cells while providing optimum dispersal and maximum recoverable counts. Aliquots of biofilm suspensions were serially diluted and plated on blood agar plates using an automated Eddy Jet Spiral Plater (IUL, SA, Barcelona, Spain). Meanwhile, propidium monoazide (PMA) combined with quantitative PCR (PMA-qPCR) was used for analysis of S. mutans cell viability as describe Klein M I et al. [Klein M I et al. 2012]. The combination of PMA and qPCR will quantify only the cells with intact membrane (i.e. viable cells) because the PMA cross-linked to DNA of dead cells and extracellular DNA modifies the DNA and inhibits the PCR amplification of the extracted DNA. Briefly, biofilm pellets were resuspended with 500 μl TE (50 mM Tris, 10 mM EDTA, pH 8.0). Using a pipette, the biofilm suspensions were transferred to 1.5 ml microcentrifuge tubes; then mixed with PMA. 1.5 μl PMA (20 mM in 20% dimethyl sulfoxide; Biotium, Hayward, Calif.) was added to the biofilm suspensions. The tubes were incubated in the dark for 5 min, at room temperature, with occasional mixing. Next, the samples were exposed to light for 3 min (600-W halogen light source). After photo-induced cross-linking, the biofilm suspensions were centrifuged (13,000 g/10 min/4° C.) and the supernatant was discarded. The pellet was resuspended with 100 μl TE, following by incubation with 10.9 μl lysozyme (100 mg/ml stock) and 5 μl mutanylysin (5 U/μl stock) (37° C./30 min). Genomic DNA was then isolated using the MasterPure DNA purification kit (Epicenter Technologies, Madison, Wis.). Ten pictograms of genomic DNA per sample and negative controls (without DNA) were amplified by MyiQ real-time PCR detection system with iQ SYBR Green supermix (Bio-Rad Laboratories Inc., CA) and S. mutans specific primer (16S rRNA) [Klein M I et al 2010].
Data are presented as the mean±standard deviation (SD). All the assays were performed in duplicate in at least two distinct experiments. Pair-wise comparisons were made between test and control using Student's t-test. The chosen level of significance for all statistical tests in present study was P<0.05.
The following examples are provided to illustrate certain embodiments of the invention. They are not intended to limit the invention in any way.
All fusion tags (CTB, PTD, protegrin, retrocyclin) were fused to the green fluorescent protein (smGFP) at N-terminus to evaluate their efficiency and specificity. Fusion constructs encoding these fusion proteins were cloned into chloroplast transformation vectors which were then used to transform plants of interest as described in U.S. patent application Ser. No. 13/101,389 which is incorporated herein by reference. To create plants expressing GFP fusion proteins, tobacco chloroplasts were transformed using biolistic particle delivery system. As seen in the
To scale up the biomass of each GFP tagged plant leaf material, each homoplasmic line was grown in a temperature- and humidity-controlled greenhouse. Fully grown mature leaves were harvested in late evenings to maximize the accumulation of GFP fusion proteins driven by light-regulated regulatory sequences. To further increase the content of the fusion proteins on a weight basis, frozen leaves were freeze-dried at −40° C. under vacuum. In addition to the concentration effect of proteins, lyophilization increased shelf life of therapeutic proteins expressed in plants more than one year at room temperature [Daniell et al 2015; 2016]. Therefore, in this study, lyophilized and powdered plant cells expressing GFP-fused tag proteins were used for oral delivery to mice.
Expression and Purification of GFP Fused Antimicrobial Peptides from Transplastomic Tobacco.
Tobacco leaves expressing GFP fused antimicrobial peptides RC101 and PG1 were harvested from greenhouse and subsequently lyophilized for protein extraction and purification. The average expression level of GFP-RC101 was found to be 8.8% of total protein in crude extracts while expression of GFP-PG1 was that of 3.8% of total protein based on densitometry. The difference in expression levels was similar to what was reported previously (Lee et al 2011, Gupta et al, 2015).
Purification of GFP fused to different antimicrobial peptides (RC101 and PG1) was done in order to test the microbicidal activity against both planktonic and biofilm forming S. mutans. Lyophilized tobacco material expressing different GFP fusions was used for extractions and subsequent downstream processing (See
We first examined the antimicrobial activity of GFP-PG1 using dose-response time-kill studies as shown in
Preventing the formation of pathogenic oral biofilms is challenging because drugs need to exert therapeutic effects following topical applications. To determine whether GFP-PG1 can disrupt the initiation of the biofilm, we treated saliva coated apatitic (sHA) surface (tooth surrogate) with a single topical treatment of GFP-PG1 for 30 min, and then incubated with actively growing S. mutans cells in cariogenic (sucrose-rich) conditions. We observed substantial impairment of biofilm formation by S. mutans with minimal accumulation of EPS-matrix on the GFP-PG1 treated sHA surface (
Disruption of Pre Formed Biofilm by AMP with or without EPS-Degrading Enzymes
Disruption of formed biofilms on surfaces is challenging. Disruption of cariogenic biofilms is particularly difficult because drugs often fail to reach clusters of pathogenic bacteria (such as S. mutans) because of the surrounding exopolysaccharides (EPS)-rich matrix that enmeshes and protects them [Bowen and Koo, 2011]. EPS-degrading enzymes such as dextranase and mutanase could help digest the matrix of cariogenic biofilms, although they are devoid of antibacterial effects. We first optimized the dextranase and/or mutanase required for EPS-matrix disruption without affecting the cell viability (data not shown). As shown in
To explore this concept, Streptococcus mutans biofilms were pre-formed on sHA surface, and treated topically with GFP-PG1 and EPS-degrading enzymes (Dex/Mut) either alone or in combination. Time-lapsed confocal imaging and quantitative computational analyses were conducted to analyze EPS-matrix degradation and live/dead bacterial cells within biofilms (
Uptake of GFP Fused with Different Tags by Human Periodontal Cells.
Purified GFP fusion proteins when incubated with human cultured cells, including HPDLS, MMS, SCC-1, GMSC, AGK and osteoblast cells (OBC) revealed interesting results. Although only one representive image of each cell line is presented, uptake studies were performed in triplicate and at least 10-15 images were recorded under confocal microscopy. Without a fusion tag, GFP did not enter any tested human cell line. Both CTB-GFP and PTD-GFP effectively penetrated all tested cell types, although their localization patterns differed. Upon incubation with CTB-GFP, GFP signals localized primarily to the periphery of HPDLSC and MMSC, uniformly small cytoplasmic puncta in SSC-1, AGK, OBC and large cytoplasmic foci in GMSC. PTD-GFP was observed as small cytoplasmic foci in MMSC, variably sized cytoplasmic puncta in HPDLSC, GMSC, AGK, OBC and both the cytoplasm and the periphery of SCC-1 cells. PG1-GFP is the most efficient tag in entering all tested human cells because GFP could be localized at tenfold lower concentrations than any other fusion proteins. PG1-GFP showed exclusively cytoplasmic localization in HPDLSC, SCC-1, GMSC and AGK cells and localized to both the periphery and cytosol in MMSC, but it is only localized to the periphery of OBC. RC101-GFP was localized in SCC-1, GMSC, AGK and OBC, but its localization in HPDLSC was negligible and was undetectable in MMSC cells.
The assembly of cariogenic oral biofilms is a prime example of how pathogenic bacteria accumulate on a surface (teeth), as an extracellular EPS matrix develops. Prevention of cariogenic biofilm formation requires disruption of bacterial accumulation on the tooth surface with a topical treatment. Chlorhexidine (CHX) is considered ‘gold standard’ for topical antimicrobial therapy (Flemmig and Beikler 2011; Marsh et al 2011; Caufield et al 2001). CHX effectively suppresses mutans streptococci levels in saliva, but it has adverse side effects including tooth staining and calculus formation, and is not recommended for daily preventive or therapeutic use (Autio-Gold 2008). As an alternative, several antimicrobial peptides (AMP) have been developed and tested against oral bacteria, and have shown potential effects against biofilms (albeit with reduced effects vs planktonic cells) (as reviewed by Silva et al., 2012) Unfortunately, most of these studies tested antibiofilm efficacy using continuous, prolonged biofilm exposure to AMPs (several hours) rather than topical treatment regimen as used clinically. Furthermore, synthetic AMPs are expensive to produce making them unaffordable for dental applications. Here, we show a plant-produced AMP, which demonstrates potent effects in controlling biofilm formation with a single, short-term topical treatment of a tooth-surrogate surface.
Developed cariogenic biofilms are characterized by bacteria embedded in EPS matrix, making biofilm treatment and removal extremely difficult (Paes Leme et al 2006; Koo et al 2013). EPS-rich matrix promotes microbial adhesion, cohesion and protection as well as hindering diffusion (Koo et al 2013; Flemming and Wingender 2010. EPS matrix creates spatial and microenvironmental heterogeneity in biofilms, modulating the growth and protection of pathogens against antimicrobials locally as well as a highly adhesive scaffold that ensures firm attachment of biofilms on tooth surfaces (Flemming and Wingender 2010; Peterson et al. 2015). CHX is far less effective against formed cariogenic biofilms (Hope and Wilson, 2004; Van Strydonck et al 2012; Xiao et al., 2012). The EPS are comprised primarily of a mixture of insoluble (with high content of α1,3 linked glucose) and soluble (mostly α1,6 linked glucose) glucans (Bowen and Koo 2011). Thus, the possibility of using EPS-matrix degrading dextranase or mutanase (from fungi) to disrupt biofilm and prevent dental caries has been explored and included in commercially available over-the-counter mouthwashes (e.g. Biotene PBF). However, topical applications of enzyme alone have generated moderate anti-biofilm/anti-caries effects clinically (Hull 1980), possibly due to lack of antibacterial action and reduced enzymatic activity in the mouth (Balakrishnan et al 2000). Interestingly, a recent in vitro study has shown that a chimeric glucanase comprised of fused dextranase and mutanase is more effective in disrupting plaque-biofilms than either enzyme alone (Jiao et al 2014). However, an approach of combining antimicrobial agents with both EPS-matrix degrading enzymes into a single therapeutic system has not yet been developed, likely due to difficulties associated with cost and formulations. In this study we demonstrate that PG1 together with matrix-degrading enzymes act synergistically and effectively to disrupt cariogenic biofilms. This feasible and efficacious topical antibiofilm approach is capable of simultaneously degrading the biofilm matrix scaffold and killing embedded bacteria using antimicrobial peptides combined with EPS-digesting enzymes.
Retention of high level antimicrobial activity by protegrin along with GFP fusion opens the door for a number of clinical applications to enhance oral health, beyond disruption of biofilms. In addition to biofilm disruption, enhancing wound healing in the gum tissues is an important clinical need. We recently reported that both protegrin and retrocyclin can enter human mast cells and induce degranulation, an important step in the wound healing process (Gupta et al 2015). Therefore, antimicrobial peptides protegrin and retrocyclin play an important role in killing bacteria in biofilms and initiate wound healing through degranulation of mast cells. In addition, it is important to effectively deliver growth hormones or other proteins to enhance cell adhesion, stimulate osteogenesis, angiogenesis, bone regeneration, differentiation of osteoblasts or endothelial cells. Previously identified cell penetrating peptides have several limitations. CTB enters all cell types via the ubiquitous GM1 receptor and this requires pentameric form of CTB. PTD on the other hand does not enter immune cells (Xiao et al 2016).
In this study we tested ability of PG1-GFP or RC101-GFP to enter periodontal and gingival cells. PG1-GFP is the most efficient tag in entering periodontal or gingival human cells because GFP signal could be detected even at ten-fold lower concentrations than any other fusion proteins. Although there were some variations in intracellular localization, PG1-GFP effectively entered HPDLSC, SCC-1, GMSC, AGK, MMSC and OBC. In contrast RC101-GFP entered SCC-1, GMSC, AGK and OBC but its localization in HPDLSC and MMSC cells were poor or undetectable. Therefore, this study has identified a novel role for protegrin in delivering drugs to osteoblasts, periodontal ligament cells, gingival epithelial cells or fibroblasts to enhance oral health. It is feasible to release protein drugs synthesized in plant cells by mechanical grinding and protein drugs bioencapsulated in lyophilized plant cells embedded in chewing gums provides an ideal mode of drug delivery for their slow and sustained release for longer duration. This overcomes a major limitation of current oral rinse formulations—short duration of contact of antimicrobials on the gum/dental surface.
Beyond topical applications, protein drugs fused with protegrin expressed in plant cells can be orally delivered to deeper layers of gum tissues in a non-invasive manner and increase patient compliance. Protein drugs bioencapsulated in plants can be stored for many years at room temperature without losing their efficacy (Su et al 2015; Daniell et al 2016). The high cost of current protein drugs is due to their production in prohibitively expensive fermenters, purification, cold transportation/storage, short shelf life and sterile delivery methods. All these challenges could be eliminated using this novel drug delivery concept to enhance oral health. Recent FDA approval of plant cells for production of protein drugs (Walsh 2014) augurs well for clinical advancement of this novel concept.
Effective treatment of biofilm-associated infections is problematic as antimicrobials often fail to reach clusters of microbes present within the surrounding extracellular matrix that enmeshes and protects them. Furthermore, development of novel therapies against biofilm-related oral diseases and maintenance of oral health needs to be cost-effective and readily accessible.
To ensure a continued supply of reagents, dextranase/mutanase and protegrin/retrocyclin are expressed independently and as fusion proteins in tobacco and other plant chloroplasts, such as lettuce. Proteins will be produced and used in low-cost purification strategies. Tobacco plants produce a million seeds, and thus, it is feasible to scale up production easily. Each acre of tobacco will produce up to 40 metric tons of biomass, facilitating low-cost, large-scale production of AMP, enzymes and fusion constructs encoding the same. In another approach, the proteins are produced in an edible plant such as lettuce.
Several dextranases (Dex) and mutanases (Mut) have been isolated from fungi and bacteria and characterized for their enzymatic activity. Optimal dextranase and mutanase enzymes should have enzymatic properties suitable for human oral environment. Based on short duration of oral treatments, strong binding/retention property to plaque-biofilms and catalytic activity to both types of EPS (dextrans and mutans) are highly desirable. The enzymes added in commercial dextranase-containing mouthwashes (e.g. Biotene) are largely derived from fungi (Penicillium sp. and Chaetomium erraticum). However, fungal dextranases show higher temperature optima (50-60° C.) than bacterial dextranases (35-40° C.). Furthermore, bacterial dextranases are more stable and effective at oral temperature (˜37° C.) and are suitable for dental caries-prevention. Recently, a dextranase from Arthrobacter sp strain Arth410 showed superior dextran degradation properties at optimal temperatures (35-45° C.) and pH values (pH 5-7) found in mouth and in cariogenic biofilms when compared to fungal dextranases. In addition, topical applications of bacterial dextranase are more effective in reducing dental caries in vivo than fungal dextranse. Likewise, a bacterial mutanase from Paenibacillus sp. strain RM1 shows that biofilm was effectively degraded by 6 hr incubation even after removal of the mutanase, preceded by first incubation with the biofilms for 3 min. Also, when compared to other microbial species, RM1 mutanase shows enhanced biofilm-degrading property. Notably, fungal enzymes require glycosylation, which precludes their expression in chloroplasts. In addition, immunogenicity of glycoproteins in human system may raise additional regulatory concerns. Therefore, the present invention involves use of bacterial dextranase and mutanase for expression in chloroplasts.
In order to increase the production of Arth410 dextranase and RM1 mutanase protein in chloroplasts, both sequences have been codon optimized for chloroplast expression. See
In order to maximize synthesis and reduce toxicity of AMPs, ten tandem repeats of PG1 or RC101, separated by protease cleavage sites as shown in
As mentioned above, the sequences encoding the AMP/biofilm degrading enzymes are optionally codon-optimized prior to insertion into chloroplast transformation vectors, such as pLD. Chloroplast transformation relies upon a double homologous recombination event. Therefore, chloroplast vectors comprise homologous regions to the chloroplast genome which flank the expression cassette encoding the heterologous proteins of interest, which facilitate insertion of the transgene cassettes into the intergenic spacer region of the chloroplast genome, without disrupting any functional genes. Although any intergenic spacer region could be used to insert transgenes, the most commonly used site of transgene integration is the transcriptionally active intergenic region between the trnI-trnA genes (in the rrn operon), located within the IR regions of the chloroplast genome (
A hydrophobic interaction column (HIC; TOSOH Butyl Toyopearl 650m) can be used to purify PG1 fused with Green Florescent Protein (GFP). The GFP selectively binds to the HIC and facilitates Rc101/PG1 to >90% purity. Despite using the expensive HIC chromatography method, recovery is very poor (<20%). To address this problem and enhance yield, 10 tandem repeats of PG1 with an elastin like biopolymer (GVGVP (SEQ ID NO: 11);
In an alternative approach, a signal peptide is fused with dextranase or mutanase for expression in E. coli, where the signal peptide will result in secretion of the enzymes into the extracellular media. In addition, secretory proteins should pass through two membrane systems of E. coli, during which they pass through the periplasmic environment where disulfide isomerases, foldases and chaperones are present. Therefore, correct folding and disulfide bond formation of secretory proteins are facilitated by the enzymes, resulting in enhancement of biological activity of proteins (ideal for AMPs). Another merit of this production strategy is the low level of proteolytic activity in the culture medium which serves to enhance the stability of the recombinant protein. The signal sequence of the secreted protein is cleaved during the export process, creating an authentic N-terminus to the native protein. There are several molecules useful for translocating proteins to extracellular media, such as TAT, SRP, or SecB-dependent pathways. However, rather than working independently, the different pathways closely interact with each other. Both SRP and SecB-dependent pathways can work together in targeting of a single protein. Also, under Sec-deficient conditions, translocation of Sec pathway substrates can be rescued by TAT systems.
Among numerous signal sequences, outer membrane protein A (OmpA) and Seq X (derived from lac Z) signal peptide demonstrate superior export functions and are capable of exporting fused protein into extracellular medium at up to 4 g/L and 1 g/L, respectively. Therefore, these signal sequences are used for efficient exporting of Arth 410 Dex and RM1 Mut to extracellular milieu. Accumulation of the dextranase and mutanase exported into media will be determined by protein quantitation and enzyme assays.
Successful expression of these proteins in E. coli has been achieved. See Western blot results shown in
The AMP-enzyme combination effectively disrupts cariogenic biofilm formation and the onset of cavitation in vivo. Furthermore, AMP-enzyme fusion protein appears to be superior to current chemical modalities for antimicrobial therapy and caries prevention.
As mentioned previously, effective AMP-enzyme (independently or in combination) can be expressed in lettuce chloroplasts under the control of endogenous lettuce regulatory elements, for large scale GLP production and stability assessment. A key advantage is the lower production cost by elimination of prohibitively expensive purification processes. Freeze-dried leaf material expressing AMP/enzymes can be stored at ambient temperatures for several months or years while maintaining their integrity and functionality. See
The steps for producing the AMP/enzymes or fusions thereof are shown in
AMP-enzyme(s) expressed in the edible plants are preferably orally delivered (topically) when used for treatment of oral diseases and the prevention and inhibition of dental carie formation. For enhanced lysis of plant cells within the oral cavity, AMP/enzyme expressing plant cells are optionally mixed with plant cells expressing cell wall degrading enzymes, described in U.S. patent application Ser. No. 12/396,382, also incorporated herein by reference.
Chewing gum tablet preparation is shown in
It is clear from these data that gum tablets comprising the AMP-enzyme fusion proteins of the invention will deliver the active material for a suitable time period to achieve bacterial kill and plaque or biofilm degradation. However, oral rinses such as Listerine® (i.e., 0.064% thymol, 0.06% methyl salicylate, 0.042% menthol, 0.092% eucalyptol, ethanol, water, benzoic acid, poloxamer 407, sodium benzoate and caramel) can also be employed to deliver the AMP-enzyme fusion proteins or combinations of the invention.
AMPS have the ability to stimulate innate immunity and wound healing, in addition to antimicrobial activity. Harnessing this novel mast cell host defense feature of AMPs in addition to their antimicrobial properties expands their clinical applications. Biofilm-associated caries is the most challenging model for development of topical therapeutics. When developed, such topical drug delivery can be easily adapted to other biofilms, as matrix formation hinders drug efficacy in many other biofilm-associated diseases. Matrix is inherent in all biofilms thus the application goes beyond the biofilm in the mouth. The biofilm inhibiting compositions described herein can also be employed in coating stents, artificial joints, implants, valves and other medical devices inserted into the human body for the treatment of disease.
As discussed above, the AMP/enzymes, or leaves expressing the same can be incorporated into a chewing gum for effective topical application of the same for the treatment of oral disease. The compositions may also be incorporated into an oral rinse, such as Listerine®. As mentioned previously, other anti dental carrie agents such as fluoride or CHX may included in such gums or oral rinses.
The references below in Table 2 describe a number of different mutanases from a variety of biological sources. Each of these references incorporated herein by reference.
Paenibacillus humicus NA1123
Paenibacillus humicus
Paenibacillus humicus
Paenibacillus sp. strain RM1
Paenibacillus sp. Strain RM 1: Identification of Its Mutan-Binding Domain, Essential for
Paenibacillus sp. strain RM1
Paenibacillus sp. Strain RM 1: Identification of Its Mutan-Binding Domain, Essential for
Streptococcus sanguinis glucans formed under various conditions. Caries Res 35: 67-
Pseudomonas sp. strain
Pseudomonas sp.
Additional biofilm degrading enzyme encoding sequences useful in the practice of the invention, include without limitation,
I) Paenibacillus humicus NA1123
See also the world wide web at ncbi.nlm.nih.gov/nuccore/AB489092 Genbank AB489092
Reference: Otsuka R, et al. Microbiol Immunol. 2015 January; 59(1):28-36.
2. The Protein Sequence of Mutanase from Paenibacillus humicus NA1123
3. Sequence of mRNA from Paenibacillus humicus NA1123
II) Paenibacillus curdlanolyticus MP-1
1. General Information of Mutanase from Paenibacillus curdlanolyticus MP-1
See the world wide web at ncbi.nlm.nih.gov/nuccore/HQ640944
Reference: Pleszczyńska M, et al. Protein Expr Purif. 2012 November; 86(1):68-74.
2. The Protein Sequence of Mutanase from Paenibacillus curdlanolyticus MP-1
3. Sequence of mRNA from Paenibacillus curdlanolyticus MP-1
III) Paenibacillus sp. Strain RM1.
AAGGPNLTPG KPITASGQSQ
3. Sequence of mRNA from Paenibacillus sp. Strain RM1
IV) Trichoderma harzianum (CCM F-470)
1. General Information of Mutanase from Trichoderma harzianum
Also see the world wide web at uniprot.org/uniprot/Q8WZM7Length: 635
Last modified: Mar. 1, 2002-v1
Checksum: iBB0D864E2F432C58
2. The Protein Sequence of Mutanase from Trichoderma harzianum
See the world wide web at uniprot.org/uniprot/Q8WZM7.fasta
3. Sequence of mRNA (Trichoderma harzianum
See the world wide web at ebi.ac.uk/ena/data/view/AJ243799&display=fasta)
(There is a polyA tail since Trichoderma harzianum is fungi)
V) Trichoderma harzianum
1. General Information of Mutanase from Trichoderma harzianum
Also see the world wide web at uniprot.org/uniprot/Q8WZM7;
2. The Protein Sequence of Mutanase from Trichoderma harzianum
See: the world wide web at uniprot.org/uniprot/Q8MZM7.fasta)
3. Sequence of mRNA (Trichoderma harzianum Further Information can be Found at the World Wide Web at ebi.ac.uk/ena/data/view/AJ243799&display=fasta)
(There is a polyA tail since Trichoderma harzianum is fungi)
Dextranase (Dex) Gene from Penicillium minioluteum
(see the world wide web at ncbi.nlm.nih.gov/nuccore/L41562.1)
The mature protein has 574 amino acids with MW at 67KD. The optimum reaction condition is pH 5.5 and 40° C. The pH range is 3-6.
2. Dextranase (Dex) Gene from Penicillium aculeatum (Talaromyees aculeatus Strain z01)
GenBank: KF999646.1 (see the world wide web at ncbi.nlm.nih.gov/nuccore/KF999646.1)
The optimum pH is around 5. The pH range is 3-6.
3. Penicillium funiculosum dexA Gene for dextranase
GenBank: AJ272066.1 (see the world wide web at ncbi.nlm.nih.gov/nuccore/7801166)
The optimum pH is around 5.5. The optimum temperature is 60° C. The pH range is 5-7.5 (see the world wide web at sciencedirect.com/science/article/pii/S0032959298001277)
The amino acid and nucleic acid sequence of the triacylglycerol lipase and other information regarding lipY can be found on the UNIPROT website on the world wide web at uniport.org/unitpro/I6Y2J4 (SEQ ID NO: 38) and on the NCBI website NCBI Reference Sequence: YP_177924.1. The nucleic acid sequence encoding lipase could optionally be codon optimized for maximal plant plastid expression using the guidance provided in
Current approaches for oral health care rely on procedures that are unaffordable to impoverished populations. As aerosolized droplets in the dental clinic and poor oral hygiene may contribute to spread of several infectious diseases, including COVID-19, new solutions for dental biofilm/plaque treatment at home are required. In this example, an affordable method for dental biofilm disruption via expression of lipase, dextranase or mutanase in chloroplast vectors in plant cells is described. The antibiotic resistance gene used to for selection of chloroplast genetransformants were subsequently removed using direct repeats flanking the aadA gene and enzymes were successfully expressed in marker-free lettuce transplastomic lines. Equivalent enzyme units of plant-derived lipase performed better than purified commercial enzymes against biofilms, specifically targeting fungal hyphae formation.
Combination of lipase with dextranase and/or mutanase suppressed biofilm development by degrading the biofilm matrix, with concomitant reduction of bacterial and fungal accumulation. In chewing gum tablets formulated with freeze-dried plant cells, expressed protein was stable up to 3 years at ambient temperature and was efficiently released in a time-dependent manner using a mechanical chewing simulator device. Development of edible plant cells expressing enzymes eliminates the need for purification and cold-chain transportation, providing a translatable therapeutic approach. Biofilm disruption through plant enzymes and chewing gum-based delivery offers an effective and affordable dental biofilm control at home particularly for populations with minimal oral care access.
A codon usage reference table for codon optimization based on codon usage frequency of psbA gene in 133 plant species was previously developed (Kwon et al., 2016). Native mutanase coding mut gene nucleotide sequence from Paenibacillus sp. was codon optimized by replacing less preferred codons with more preferred ones, which eventually generated codon usage frequency close to that in reference psbA gene. Rare codons in the native mut gene sequence with a frequency <5% in the reference were replaced by more preferred codons (
PG1-Smdex and Mut Gene (Co) Cloning in pLsLF-Marker-Free Chloroplast Vector
The native Smdex gene sequence from S. mutans (ATCC 25175) fused with PG1 (Protegrin-1 encoding) downstream, and codon optimized mut gene were synthesized (GenScript Biotech, Piscataway Township, NJ) (Liu et al., 2016). The PG1-Smdex and synthesized mut (co) genes were cloned into pLsLF-marker-free vector using NdeI and PshAI restriction sites and transformed into TOP10 E. coli cells. The lettuce chloroplast transformation vector pLsLF, which contains spectinomycin-resistant gene (aadA, aminoglycoside 3′-adenylytransferase gene) as selectable marker, was used as a backbone.
To design a marker-removable vector, the 649-bp long direct repeat DNA sequence, derived from atpB promoter and 5′ UTR (Daniell et al., 2019a,b; Kumari et al., 2019), was PCR amplified using lettuce total genomic DNA as a template. The sequence-confirmed direct repeats were then cloned to flank aadA expression cassette. For the insertion of single-digested atpB fragments into the vector backbone, NEBuilder HiFi DNA (NEB, Ipswich, Mass.) assembly kit was used to avoid the possible ligation of the fragments in a reverse direction.
The successful insertions were confirmed by restriction digestion. Expression of PG1-dextranase and mutanase in E. coli was confirmed by Western blot. The pLsLF-MF-PG1-dextranase and pLsLF-MF-mutanase (co) plasmid were extracted using PureYield™ plasmid Midiprep System (Promega, Madison, Wis.) and used for subsequent particle bombardment.
The pLsLF-MF-PG1-dextranase and pLsLF-MF-mutanase (co) plasmids were transformed into 1-month-old lettuce (L. sativa) leaves by bombardment as previously described (Lee et al., 2011). After the bombardment, lettuce leaves were cut into small pieces (<1 cm2) and grown on regeneration media containing spectinomycin (50 mg/mL) as described previously (Daniell et al., 2019a,b; Kumari et al., 2019; Lee et al., 2011; Ruhlman et al., 2010). The integration of pLsLF-MF-PG1-dextranase and pLsLF-MF-mutanase (co) vectors in regenerated shoots were validated by Southern blot and PCR using specific primers sets:
The PCR positive leaves were subjected to the second round of selection on regeneration media containing spectinomycin (50 mg/L). Any regenerated shoots showing bleached leaves were immediately evaluated by PCR analysis to confirm the excision of aadA gene and were then transferred to spectinomycin-free rooting media to induce roots. Once the roots were formed, homoplasy was confirmed by Southern blots as described below. Expression of enzymes in homoplasmic lines were confirmed using Southern blots and enzyme assays as described previously (Daniell et al., 2019a,b; Kumari et al., 2019; Ruhlman et al., 2010; Verma et al., 2008).
For Southern blotting, 2 μg total genomic DNA from untransformed WT, T1 and T2 generation marker-free plants integrated with PG1-Smdex, lipY and T0 generation integrated with mut (co) were digested by suitable restriction enzymes and separated in 0.8% agarose gel, transferred onto the nylon membranes (Nytran, GE Healthcare), and probed as described previously (Kumari et al., 2019; Kwon et al., 2018).
Seeds from untransformed WT and previously developed three independent T0 transplastomic lipase (Kumari et al., 2019) expressing plants were germinated on ½ MS medium without any antibiotics. The germinated seedlings were transferred and grown in magenta box. The genomic DNA from leaves of two different T1 plants in each of the three independent events (in total six plants) and WT was isolated, and digested with SmaI restriction enzyme (New England Biolabs, Hertfordshire, UK).
Leaves from marker-free transplastomic plants expressing lipase, PG1-dextranase, mutanase and untransformed WT plants were stored at −80° C. and freeze-dried in a lyophilizer as described previously (Daniell et al., 2019a,b). Lyophilized leaves were ground into fine powder in a blender and used as the source material for the proteins/enzyme extraction. Total soluble proteins (TSP) was extracted by suspending 50 mg of plant powder in 1 ml plant extraction buffer (respective buffer of each enzymes and EDTA-free protease inhibitor cocktail (Thermo Scientific, Waltham, Mass., USA) and kept on a mixer (Eppendorf) at 4° C. for 1 h. Samples were sonicated, centrifuged at 9391 g for 30 min and the supernatant (TSP) was collected. TSP was quantified by Bio-Rad protein assay dye (Bio-Rad, Hercules, Calif., USA) by following the manufacturer's protocol. Bovine serum albumin (BSA) protein was used as standard. The impact of sonication during extraction of TSP and the role of PIC in the extraction buffer was evaluated by independent experiments. The stability of recombinant enzyme in each crude extract was evaluated by activity assay and compared.
Leaves harvested at two different time points (30 and 45 days) from two independent marker-free PG1-dextranase transplastomic events (Plant-46 and Plant-47) were lyophilized, ground into powder and evaluated for the enzyme activity. Blue dextran plate assay was performed for qualitative enzyme activity analysis. Blue dextran substrate (from Leuconostoc mesenteroides, Sigma, Louise, Mo., USA) and agar were added into 100 mM sodium acetate buffer (pH 5.5) at final concentration of 0.5% and 1.25% respectively. The suspension was boiled, mixed properly and poured into the plate. After solidification, small wells were created and 50 μg of plant crude extract (TSP) from PG1-dextranase transplastomic plants and WT plant (as negative control) was loaded into the wells. The plate was incubated at 37° C. overnight. Enzymatic activity of PG1-dextranase was visualized in the form of a halo caused by the breakdown of blue dextran around the well. Purified dextranase enzyme from Penicillium sp. (Sigma) was used as positive control.
An enzyme assay was performed to quantify dextranase activity in the crude extract of transplastomic PG1-dextranase plants. TSP of PG1-dextranase transplastomic and WT plants (50 μL) was incubated with 50 μL of dextranase (1% in 100 mm sodium acetate pH 5.5), incubated at 37° C. for 1 h and the released sugar was estimated by dinitrosilycilic acid method (Kumari et al., 2019) using maltose as standards. The enzymatic assay was performed in triplicates and the data are presented as mean and standard deviation. The enzyme activity was represented as sugar released μmol/h/g dry weight of plant powder. The importance of sonication for the release of PG1-dextranase enzyme from the plant powder and PIC for the stability of the released enzyme in the crude plant extract was also evaluated. Plant powder of 45 days old leaves from plant-46 was used for these experiments.
Lipase enzyme assay was performed using the method described previously (Kumari et al., 2019). Briefly, TSP was extracted from the lyophilized plant powder in 100 mM sodium phosphate buffer (pH 8.0) and was quantified. In the assay, 50 μL of TSP and 100 mM p-nitrophenyl butyrate (5 μL) was mixed in 450 μL reaction buffer (100 mM sodium phosphate buffer pH 8 and 0.9% NaCl) and incubated at 37° C. for 10 min. Released p-nitrophenol (hydrolysed product of p-nitrophenyl butyrate) was estimated by using different known concentrations of p-nitrophenol as standards. The enzyme assay was performed in triplicate and data are presented as mean and standard deviation. The enzyme activity was represented as p-nitrophenol released μmol/h/g dry weight. The effect of sonication during the protein extraction and stability of enzyme in the absence of PIC was also examined for plant-derived lipase as described earlier.
Lyophilized plant powder was extracted with a ratio of 50 mg powder/1 mL extraction buffer [0.1M sodium acetate buffer (pH 5.5)], followed by vortex homogenization at 4° C. for 1 h (Eppendorf 5432) and sonication for 3 cycles (5 s on, 10 s off, 80% amplitude). The homogenate was centrifuged at 9391 g for 30 min and the supernatant was transfer into a new tube. Then the supernatant was dialysed as follows: 10 mL of plant protein extraction was sealed in 15 cm of semi-permeable membrane (MWCO: 20 KD) and placed in 2 L of extraction buffer for 4 h. This buffer was replaced with fresh buffer and incubated at 4° C. overnight (16 h). The total protein concentration was determined by Bradford assay.
The activity of mutanase in the dialysed plant crude was determined by enzymatic assays as described with some modifications (Kopec et al., 1997; Verma et al., 2008). Crude extracts of transplastomic plants were incubated with the substrate (mutan or α-1, 3-linked glucans) in 0.1 M sodium acetate buffer (pH 5.5) at 37° C. for 60 min. The amount of reducing sugar released was determined by the Somogyi-Nelson method (Somogyi, 1945). Crude extracts of untransformed plant (WT) with equal protein concentrations were used to check the endogenous mutanase activity from plant cells. One unit (U) of mutanase activity was defined as the amount of enzyme that releases 1.0 μmol of reducing sugar from mutan per hour at pH 5.5 at 37° C. Mutanase (EC3.2.1.59) purified from Bacillus sp. fermentation (Amano Enzyme, Japan) was used as the commercial enzyme standard.
Chewing gum tablets containing ground plant powder were prepared by compression process. Gum tablets contained the gum base (27.71%), sorbitol (17.18%), maltitol (14.78%), xylitol (13.86%), isomalt (10.07%), natural and artificial flavors (7.21%), magnesium stearate (2.95%), silicon dioxide (0.82%), stevia (0.42%) and plant cell powder (5%) in order to offer the best flavor, taste, softness and compression. The gum tablet chews and performs exactly like the conventional chewing gum based on physical characteristics.
For stability assay, 125 mg of gum tablet was ground with 500 μL protein extraction buffer [(0.2 M Tris HCl pH 8.0, 0.1 M NaCl, 0.01 M EDTA, 0.4 M sucrose, 0.2% Triton X supplemented with 2% PMSF and a PIC (Pierce, Waltham, Mass., USA)], followed by sonication at 80% amplitude for 5 s for two times. After sonication, the samples were centrifuged at 13 523 g at 4° C. for 10 min. Then 100 μL of supernatant was loaded to a fluorescent microplate reader where the GFP was detected at 485 nm (excitation) and 538 nm (emission) using the commercial GFP (Vector Laboratories, San Diego, Calif., USA) as a standard. Release of GFP from gum tablets was studied using a Universal Mechanical Testing Machine equipped with Merlin software (Instron Model 5564, Norwood, Mass.) in cyclic loading mode.
Chewing gum tablets (25 mg) were placed in 10 mL of artificial saliva (Pickering Laboratories, Mountain view, CA) in a polycarbonate chamber and loaded cyclically in compression using a piston attached to a load cell. A load range of −1.5 to −500N for intervals of 1, 5, 7 and 10 min and cycles of 55, 287, 364 and 591 to simulate human chewing. A wide range of bite forces have been reported for adult humans. Values ranging from 1300N to 285N have been reported (Takaki et al., 2014; Yong, 2010). Varga et al. (2011) reported mean bite force values of 522N for males and 465N for females with normal occlusion. A representative compressive load of 500N was selected for this study. The GFP concentrations in both the supernatant and the pellet after 1, 5, 7, and 10 min were determined by the same above-mentioned method.
Candida albicans SC5314, a well-characterized fungal strain and Streptococcus mutans UA159 serotype c (ATCC 700610), a cariogenic dental pathogen and well-characterized EPS producer were used in cross-kingdom biofilm experiments. To prepare the inoculum used in this study, C. albicans (yeast form) and S. mutans cells were grown to mid-exponential phase in ultrafiltered (10-kDa molecular-mass cutoff membrane; Millipore, MA) tryptone-yeast extract broth (UFTYE; 2.5% tryptone and 1.5% yeast extract) with 1% (wt/vol) glucose at 37° C. and 5% CO2 as described previously (Falsetta et al., 2014).
Biofilms were formed using saliva-coated hydroxyapatite (sHA) disc model as described above and by Falsetta et al., 2014 and Hwang et al., 2017. Briefly, the hydroxyapatite discs (surface area, 2.7±0.2 cm2; Clarkson Chromatography Products, Inc., South Williamsport, Pa.) coated with filter-sterilized, clarified whole saliva were vertically suspended in a 24-well plate using a custom-made disc holder (
Biofilms were maintained at 37° C. under 5% CO2. To access the antibiofilm efficacy of the plant crude extracts, we developed a topical treatment regimen for feasible clinical applications (
Biofilms were grown until 19 h and were subject to microbiological analyses, including the total number of viable cells (CFU) of bacteria/fungi and the total biomass on each sHA disk (dry weight) as detailed elsewhere (Hwang et al., 2017). Briefly, at 19 h, the biofilms were harvested from the sHA disks and homogenized via optimized sonication procedure, which does not kill fungal or bacterial cells while providing maximum recoverable counts (Koo et al., 2013). Aliquots of biofilm suspensions were serially diluted and plated on blood agar plates and the plates were grown at 37° C. under 5% CO2 for 2 days. Bacterial and fungal viability in the biofilm was accessed by determining their respective CFU recovered on the blood agar plates. The amount of biofilm dry weight (biomass) was also determined.
The impact of the topical treatments was assessed by examining the 3D architecture and the spatial distribution of Gtf-derived EPS glucans and fungal/bacterial cells within live biofilms using well-established protocols optimized for biofilm imaging and quantification (Falsetta et al., 2014; Hwang et al., 2017). Briefly, the EPS glucan matrix was labelled via incorporation of AlexaFluor 647 dextran conjugate (Molecular Probes Inc., Eugene, Oreg.) throughout the biofilm formation.
S. mutans was stained with Syto 9 (Molecular Probes), while C. albicans cell wall was labeled with Concanavalin A-tetramethyl rhodamine conjugate (Molecular Probes). High-resolution confocal imaging was performed using confocal laser scanning fluorescence microscope (LSM800 with Airyscan, Zeiss, Germany) equipped with a 20×(1.0 numerical aperture) water immersion objective. Each biofilm was scanned at five randomly selected areas, and confocal image series were generated by optical sectioning at each of these positions. Computational analysis of confocal images using the advanced biofilm 3-dimensional analysis tool BiofilmQ (available on the world wide web at: drescherlab.org/data/biofilmQ) was conducted to determine the biovolumes of bacteria, fungi and EPS in order to complement our microbiological analysis (Hartmann et al., 2021). ImageJ (FIJI) was used for post-acquisition image processing and creating 3-dimensional renderings of biofilm architecture (Schindelin et al., 2012).
To investigate the impact of the treatments on bacterial/fungal cell viability within the mixed biofilm, in situ cell viability staining was performed and followed by detailed imaging at single-cell resolution using cell membrane integrity as a biomarker for viable cells. TOTO-3 (Molecular Probes), a cell impermeable dimeric cyanine acid dye as a dead cell indicator for both bacterial and fungal cells because of its high affinity for nucleic acids, was used (Chiaraviglio and Kirby, 2014). Thus, when the microbial cells are killed and the plasma membrane integrity are compromised, these probes will enter cells, bind to nucleic acids, and exhibit a strong fluorescence.
Biofilms were stained using the optimized 1μM of TOTO-3 in 0.9% sodium chloride at 37° C. for 10 min and were counterstained with 0.65μM SYTO9 (a cell-permeable dye). Concanavalin A-tetramethylrhodamine conjugate was used to label fungal cell wall as described previously (Falsetta et al., 2014). The biofilms were sequentially scanned (488/640 nm lasers for SYTO9/TOTO-3, then 561 nm laser for Concanavalin A-tetramethylrhodamine) and the fluorescence emitted was collected using optimum emission wavelength filters (Zeiss LSM800 confocal microscope with Airyscan). Each biofilm was scanned at least three randomly selected areas. ImageJ FIJI was used for image processing and to create representative multi-channel images (Schindelin et al., 2012).
The Smdex gene (2574 bp, gene designation from Kim et al., 2011) encoding dextranase was isolated from S. mutans strain ATCC 25175 genomic DNA using PCR (SEQ ID NOs: 34 and 35) and fused to the PG1 (encoding antimicrobial peptide Protegrin-1). The mut gene (3780 bp, gene designation from Otsuka et al., 2015) of Paenibacillus sp. encoding mutanase was codon optimized to improve its translation efficiency in plant chloroplasts based on psbA genes from 133 plant species as described previously (Kwon et al., 2016). Within total 1260 codons of mut gene, 576 codons including 327 rare codons were replaced by more highly preferred codons, resulting in an increased AT content from 44% to 57% (SEQ ID NOs: 36 and 37).
The Smdex, mut and lipY (encoding lipase, gene designation from Deb et al., 2006) genes (native or codon optimized) were cloned into the newly designed marker-free chloroplast vector pLsLF-MF as described previously (Daniell et al., 2019a,b; Daniell et al., 2020; Kumari et al., 2019; Park et al., 2020) and the constructed plasmids were then delivered by gene gun into lettuce (Lactuca sativa) cv. Simpson Elite leaves (Ruhlman et al., 2010). The successful Marker-free events were identified by screening shoots for presence of the transgene cassette but absence of the antibiotic resistance gene aadA by PCR using primers described above (
To characterize homoplasmic status of transplastomic lines, total plant gDNA was extracted from marker-free Protegrin-dextranase transplastomic plants, digested with HindIII, and probed with the DIG-labeled trnI and trnA flanking sequence (
Most importantly, homoplasmic marker-free PG1-Smdex cassette was stably maintained in T1 and T2 generations of 46-1 and 46-2 lines in the absence of the antibiotic resistance gene (
The stability and inheritance of lipY gene in T1 generation and its homoplasmic status was confirmed. The single hybridizing fragment of 5.6 kb in the transplastomic and 3.13 kb in untransformed WT plants after SmaI restriction digestion of gDNA was detected in Southern blot when probed with trnI/trnA genes, flanking the expression cassette (
On the plate assay, protein crude extracts from all four tested leaf harvests (30 and 45 days of P-46 and P-47) and the purified commercial dextranase from Penicillium sp. (positive control) produced halo rings on blue dextran, while no halo formation was observed from untransformed WT plant extracts (
Enzyme release from freeze-dried plant cells with or without sonication, showed similar enzyme activity from both preparations (
Lipase activities in matured leaves of the transplastomic line and untransformed WT were 12 542.52±257.03 and 522.76±12.85 μmol/h/g dry weight, respectively with sonication and PIC in the extraction buffer. Lipase extracted without sonication but with PIC showed almost the same level of activity (
Efficacy of the plant-derived lipase was evaluated by employing a mixed-kingdom biofilm model and a treatment regimen based on topical exposure (
Quantitative computational analysis was also performed using the images acquired via confocal microscopy. The plant-lipase crude extract significantly reduced the total biovolume of the mixed biofilm (
To investigate whether the plant-derived glucanohydrolases could be used as antibiofilm therapeutics, the bioassay was performed using a mixture of dextranase and mutanase (5:1 activity ratio) that was optimized previously using purified enzymes to provide maximum matrix-degrading activity (Ren et al., 2019). Commercial purified dextranase and mutanase (at the same 5:1 ratio) was used as positive-control. The data showed that both plant-dextranase/mutanase extract and the equivalent purified enzymes were highly effective in disrupting EPS glucans, resulting in near abrogation of glucan-matrix in the mixed-kingdom biofilm (
Further computational analysis confirmed the inhibitory effects exerted by glucanohydrolases. As expected, the EPS was effectively degraded compared to vehicle control (>90% reduction, P<0.01;
To develop a therapeutic solution for fungal-bacterial mixed biofilms, a combinatorial approach using dextranase/mutanase and lipase to enhance the antibiofilm efficacy was employed. The data show that the topical treatment with dextranase/mutanase and lipase is remarkably effective, resulting in near-complete suppression of mixed-biofilm formation (
To further investigate the impact of the treatment on the biofilm accumulation, the Total Biofilm Inhibition (TBI) index (
The feasibility of incorporating the plant-derived proteins in chewing gum was tested as an alternative, easy-to-use and more affordable delivery approach (
There was no detectable loss of protein in gum tablets preparation process based on GFP quantification (
Release of GFP from gum tablets was studied using a Universal Mechanical Testing Machine by placing chewing gum pellets in 10 mL of artificial saliva in a polycarbonate chamber and loaded cyclically in compression using a piston attached to a load cell. GFP-protegrin concentration in saliva increased from 225 μg/mL in 1 min to 809 μg/mL in 10 min in the supernatant and decreased from 988 μg/mL at 1 min in the pellet to 502 μg/mL at 10 min, confirming steady release during the chewing process (
Quantification of GFP in gum tablets showed that GFP was stable in gum tablets when stored at ambient temperature for 3 years (
In order to address the high cost of production and delivery of recombinant therapeutic proteins, protein drugs (PDs) have been expressed in plant chloroplasts and orally deliver them through bioencapsulation within plant cells (Daniell et al., 2016; Daniell et al., 2021; Daniell et al., 2019a,b). Thin lettuce leaves facilitate rapid removal of water through lyophilization and offer an ideal system for expression and delivery of PDs. This platform has advanced to deliver therapeutic proteins in the clinic, yet it remains unexplored in dental medicine. Mechanical disruption, using manual or electric toothbrushes, can remove dental plaque, but they are cumbersome to use, have low compliance, and are costly. Furthermore, current antimicrobial agents to treat cariogenic plaque are inefficient due to the presence of the EPS matrix (Autio-Gold, 2008; Koo et al., 2017; Ren et al., 2019).
Matrix-degrading enzymes can effectively target the biofilm structure and enhance antimicrobial killing, while simultaneously weakening its mechanical stability, thereby promoting bacterial removal (Autio-Gold, 2008; Koo et al., 2017; Ren et al., 2019). However, the development of clinically feasible approaches is hindered by high costs for mass production as the enzymes require complex microbial fermentation and expensive purification procedures.
Here, we describe the expression of mutanase, dextranase and lipase in plant cells and synergistic efficacy when combined with plant-derived lipase for oral biofilm prevention. Although microbial dextranase has been extensively used in food industry with demonstrated safety (Purushe et al., 2012), dextranase has never been expressed in plant cells. Importantly, creation of marker-free edible plant cells expressing enzymes with high yield and stability eliminates the need for purification, providing a translatable therapeutic approach.
Current chemical modalities to treat biofilm-associated infections are primarily targeting individual bacterial or fungal components despite the importance of the extracellular EPS matrix in biofilm antimicrobial tolerance (Koo et al., 2017). In addition, targeting EPS can also disrupt the viscoelastic properties of biofilms and further weaken its mechanical stability, promoting bacterial dispersal or removal (Ren et al., 2019). The data presented herein shows that the lipase, dextranase and mutanase crude plant extracts have potent antibiofilm efficacy that can replace the costly purified enzyme standards of equivalent enzyme units. Although lipase has been approved by FDA to treat several metabolic disorders it remains underexplored to treat dental plaque.
In the present study, lipase remarkably reduced fungal load within treated biofilm by abrogating hyphal formation. This a crucial step in cross-kingdom biofilm development as hyphae provide a scaffold for co-species adhesion and EPS accumulation (Prabhawathi et al., 2014). The precise role of lipids in biofilm formation remains unclear, but recent data suggest that lipids appear to be important membrane components modulating fungal morphogenesis and hyphal elongation (Rella et al., 2016). Hence, the data presented herein indicates that lipase treatment inhibits the filamentous Candida growth impacting cross-kingdom biofilm scaffolding and accumulation. Additionally, plant-derived dextranase and mutanase degraded EPS glucan effectively inhibiting the matrix development and bacterial accumulation. However, after treatment by dextranse and mutanase, fungal cells were unaffected and detectable EPS was still present.
Interestingly, lipase has been shown to have antimicrobial properties possibly through degradation of the membrane lipids (Prabhawathi et al., 2014; Seghal Kiran et al., 2014). Using high-resolution confocal microscopy, we found that the combination of glucanohydrolases and lipase displayed enhanced antimicrobial activity against both species in the biofilms. The data also revealed that degradation of EPS glucans by dextranase and mutanase can locally expose the embedded S. mutans and C. albicans cells within the treated biofilm. This indicates that dextranase and mutanse provide access for lipase to the fungal/bacterial cell surface thereby causing microbial death. Thus, the three-enzyme combination treatment suppressed the pathogenic fungal-bacterial biofilm formation via a complementary mechanism between the effective EPS degradation by glucanohydrolases and the antimicrobial effects of lipase in situ, thereby potentiating antibiofilm efficacy.
The use of chewing gum supplemented with antimicrobial and antibiofilm therapeutics to improve treatments of dental diseases has been proposed as a promising alternative in oral health care applications (Wessel et al., 2016). Yet, no such products are currently available due to high costs for inclusion of therapeutic additives and practical formulation development.
Data presented herein indicate that the combination of dextranase and mutanase at specific amounts and ratios (as employed here) is required to maximize EPS degradation and achieve biofilm disruption. We also demonstrate that the addition of lipase-mediated antimicrobial effects to the dextranase and mutanase combinations resulted in potentiation of effective biofilm eradication.
Here, we demonstrated the feasibility of using chewing gum to release plant-derived proteins in saliva solution following mechanical bite forces. Furthermore, the proteins were stable in chewing gum stored at ambient temperature (up to 3 years), indicating a practical and easy-to-use topical delivery platform. Altogether, we provide a conceptual framework for plant made biofilm-degrading enzymes and chewing gum-based protein delivery as an innovative and affordable approach for controlling dental biofilm. This strategy helps address the societal issue of inequity in access dental care services by enabling a low-cost technology for improvement of oral health, which is also relevant during the current global crisis caused by COVID-19.
Streptococcus mutans strain ATCC 25175
While certain of the preferred embodiments of the present invention have been described and specifically exemplified above, it is not intended that the invention be limited to such embodiments. Various modifications may be made thereto without departing from the scope and spirit of the present invention, as set forth in the following claims.
This application is a Continuation-in-part of U.S. application Ser. No. 16/301,023, filed Nov. 13, 2018, which is a § 371 of International Application No. PCT/US17/32437, filed May 12, 2017, which claims priority to U.S. Provisional Application No. 62/335,650 filed May 12, 2016, the entire disclosure of each of the foregoing applications being incorporated herein by reference as though set forth in full.
This invention was made with government support under Grant Nos: R01 HL107904 and R01 HL109442 awarded by the National Institutes of Health. The government has certain rights in the invention.
| Number | Date | Country | |
|---|---|---|---|
| 62335650 | May 2016 | US |
| Number | Date | Country | |
|---|---|---|---|
| Parent | 16301023 | Nov 2018 | US |
| Child | 17672438 | US |
