ABSTRACT
The cell wall of Gram-positive bacteria contains abundant surface-exposed carbohydrate structures that are highly conserved. While these properties make surface carbohydrates ideal targets for immunotherapy, carbohydrates elicit a poor immune response that results primarily in low-affinity IgM antibodies. In a previous publication, we introduced the lysibody approach to address this shortcoming. Lysibodies are engineered molecules that combine a high-affinity carbohydrate-binding domain of bacterial or bacteriophage origin and an Fc effector portion of a human IgG antibody, thus directing effective immunity to conserved bacterial surface carbohydrates. Here, we describe the first example of a lysibody containing the binding domain from a bacteriocin, lysostaphin. We also describe the creation of five lysibodies with binding domains derived from phage lysins, directed against Staphylococcus aureus. The lysostaphin and LysK lysibodies showed the most promise and were further characterized. Both lysibodies bound a range of clinically important staphylococcal strains, fixed complement on the staphylococcal surface, and induced phagocytosis of S. aureus by macrophages and human neutrophils. The lysostaphin lysibody had superior in vitro activity compared to that of the LysK lysibody, as well as that of the previously characterized ClyS lysibody, and it effectively protected mice in a kidney abscess/bacteremia model. These results further demonstrate that the lysibody approach is a reproducible means of creating antibacterial antibodies that cannot be produced by conventional means. Lysibodies therefore are a promising solution for opsonic antibodies that may be used passively to both treat and prevent infection by drug-resistant pathogens.
INTRODUCTION
Staphylococcus aureus is a major human pathogen that is a common cause of skin and soft tissue infections (SSTI), including impetigo, folliculitis, furuncles, and subcutaneous abscesses. It also causes a range of invasive infections, such as septic arthritis, osteomyelitis, septicemia, endocarditis, meningitis, pneumonia, and biofilm formation on prosthetic devices (1, 2). There are approximately 80,000 invasive infections per year in the United States, leading to 11,000 deaths (3–7). Antibiotic resistance among clinical S. aureus isolates is a growing concern. As much as 60% of hospital-acquired S. aureus infections involve methicillin-resistant Staphylococcus aureus (MRSA), and resistant strains are now also common among community acquired infections (2, 8–10). MRSA infections have a worse prognosis and increased hospitalization costs compared to infections caused by antibiotic-sensitive strains (11, 12). Moreover, resistance to standard-of-care antibiotics for MRSA infections has been documented, including resistance to vancomycin (13), daptomycin (14), and linezolid (15). As such, it is clear that new therapeutic solutions are required to address this pathogen.
Therapeutic antibodies represent a promising treatment alternative for antibiotic-resistant pathogens (16, 17). In particular, substantial efforts have been directed toward the development of vaccines and therapeutic antibodies against S. aureus, primarily targeting surface-attached or secreted virulence factors. However, none of these have led to an approved therapeutic solution to date (18–21). Cell wall carbohydrates are attractive alternative targets for the development of therapeutic antibodies and have several advantages compared to more traditional protein targets. Wall carbohydrates are major components of the bacterial cell wall, comprising up to 60% of its dry weight, and have important roles in maintaining the proper structure and function of the bacterial cell envelope. Certain carbohydrates are covalently attached to the peptidoglycan, while others are attached to the plasma membrane by a lipid anchor; both types are often exposed on the bacterial surface. Additionally, these molecules are highly conserved within species and often at the genus level (22–28). Nevertheless, production of conventional therapeutic antibodies for carbohydrates is complicated by their very low immunogenicity. Unlike proteins, carbohydrates are not naturally processed for presentation on major histocompatibility complexes (MHC) and are therefore T cell-independent antigens. As a result, the immunological response to carbohydrates is characterized by the production of low-affinity IgM antibodies and by the absence of class switch and memory. One approach to address this is by conjugating specific carbohydrates to carrier proteins to be used in immunization. While this approach has been successful in certain cases, creation of effective opsonic antibodies against carbohydrates still represents a substantial challenge (29, 30).
In our previous publication, we introduced lysibody technology as an alternative approach to create effective opsonic antibodies against bacterial surface carbohydrates (31). Lysibodies are a chimera between the Fc region of a human IgG antibody and a carbohydrate-binding domain derived from a cell wall hydrolase. Bacteriophages produce cell wall hydrolases during their lytic cycle, termed lysins, to degrade the cell wall of an infected bacterial host and release progeny phage (32). Bacteria use cell wall hydrolases termed autolysins to remodel their cell wall during division and elongation. We demonstrated that functional lysibodies could be produced with a carbohydrate binding domain of either bacterial or bacteriophage origins, and that the binding domain can be fused at the N terminus of the Fc molecule (directly replacing the Fab region), or at the C terminus (31). Lysibodies had all the effector functions of normal IgG antibodies, demonstrated robust complement fixation and induction of phagocytosis, and were able to protect mice from a MRSA lethal challenge (31).
In this work we demonstrate for the first time that lysibodies can be created using the binding domain from a bacteriocin, lysostaphin. We also created five additional lysibodies with binding domains derived from phage lysins. These lysibodies bound to the cell wall of S. aureus to various degrees, which directly correlated with their ability to promote uptake by phagocytes. Lysostaphin and LysK lysibodies had the best activity and were thus characterized in detail. These lysibodies induced complement deposition on the surface of S. aureus and promoted the phagocytosis of staphylococci by macrophages and neutrophils. In particular, the lysostaphin lysibody showed robust in vitro activity that was superior to that of previously characterized lysibodies and efficiently protected mice in a bacteremia/kidney abscess MRSA infection model.
RESULTS
Construction of S. aureus-specific lysibodies.The aims of the current study were to construct and characterize additional lysibodies specific for S. aureus and to define the best candidate for clinical development. For this purpose, we constructed lysibodies with the binding domains of lysostaphin, a bacteriocin produced by Staphylococcus simulans biovar staphylolyticus (33), the S. aureus-specific lysin LysK (34), and four putative lysin binding domains from various sequenced staphylococcal genomes (see the supplemental material). The general design of these lysibodies is similar to that of the previously described ClyS lysibody (Fig. 1A and B) (31). We named the resulting lysibodies lysostaphin lysibody, LysK lysibody, PlySa4 lysibody, PlySa6 lysibody, PlySa7 lysibody, and PlySa32 lysibody. Lysibodies were expressed in 293T cells to allow correct glycosylation and purified by metal affinity chromatography. Purified lysibodies were run on an SDS-PAGE in the presence or absence of β-mercaptoethanol (BME), which breaks the disulfide bonds between the subunits of the lysibody homodimer. In the presence of BME all lysibodies displayed bands of a molecular weight compatible with a monomer, while in the absence of BME, lysibodies displayed a band with the molecular weight of a dimer (Fig. 1C). This demonstrates that all of the lysibodies preferentially formed homodimers stabilized by disulfide bonds, resembling the structure of an IgG antibody without light chains. In some cases, additional bands were seen, and these likely represent degradation products; only minimal secondary bands were seen for the lysostaphin lysibody, the main lysibody developed in this study.
Design and production of lysibodies. (A) Schematic representation of lysibody structure. (B) Structure of the expression vector for lysibodies. (C) Lysibodies were separated by 10% SDS-PAGE and examined by either Coomassie staining or Western blotting, using anti-human IgG horseradish peroxidase conjugate. Samples were loaded in duplicates, either with or without β-mercaptoethanol (BME).
Identification of the best lysibody candidates for further characterization.As a first step in determining the functionality of the lysibodies, we tested their ability to bind the cell wall of a protein A-negative S. aureus strain, Wood 46, using fluorescence microcopy (Fig. 2). All six lysibodies bound the cell wall of this strain, while controls showed no binding. While a protein A-negative strain was used for these microscopy experiments to avoid a nonspecific signal, in a previous study we showed that protein A is saturated with a large amount of nonspecific antibodies present in human serum (31) (similar to observations made for Streptococcus pyogenes M protein [35]), enabling lysibody activity in this environment. We then used a modified enzyme-linked immunosorbent assay (ELISA) to compare the binding of different lysibodies to S. aureus in a quantitative fashion. In this assay, the protein A-negative strain Wood 46 was immobilized on the bottom of a 96-well plate, allowing quantification of lysibodies binding to their native target. The lysostaphin lysibody showed the best binding, followed by LysK, PlySa7, and PlySa4 lysibodies (Fig. 3A). PlySa32 and PlySa6 lysibodies showed only minor binding in this assay. Given the poor binding of PlySa6 lysibody and its very low yield following purification, this lysibody was not characterized further.
Lysibodies bind to S. aureus cell wall. Log-phase S. aureus Wood 46 (protein A negative) were fixed, attached to glass cover slides, and blocked. Lysibody binding was determined by immunofluorescence microscopy using anti-human IgG Fc Alexa Fluor 594 conjugate. Bar, 5 μm.
Preliminary functional characterization of lysibodies. (A) ELISAs to evaluate the binding of different lysibodies to S. aureus were conducted by immobilizing S. aureus strain Wood 46 (protein A negative) at the bottom of a 96-well plate. Wells were incubated with serially diluted lysibodies, followed by anti-human Fc alkaline phosphatase conjugate. (B) Raw 264.7 macrophages were incubated with S. aureus Newman/pCN57 (green fluorescent protein [GFP]) in the presence of lysibodies at different concentration. Percent phagocytosis was determined by flow cytometry. Experiments were done in duplicates. (C) HL-60 neutrophils were incubated with fluorescein isothiocyanate (FITC)-labeled S. aureus Wood 46 in the presence of serially diluted lysibodies, and 0.5% complement. Experiments were done in triplicates. Percent phagocytosis was determined by flow cytometry. Error bars represent standard deviation; dashed lines represent phagocytosis level in the presence of PBS alone.
We next performed preliminary in vitro characterization of the abilities of different lysibodies to induce phagocytosis of staphylococci by Raw 264.7 macrophages (Fig. 3B) and HL-60 neutrophils (Fig. 3C) using flow cytometry. In these assays, the lysostaphin lysibody performed the best, followed by LysK, PlySa7, and PlySa4 lysibodies, whereas PlySa32 had little to no activity. Thus, there was a high degree of correlation between the level of binding of different lysibodies to S. aureus, as determined by the ELISA, and their ability to induce phagocytosis. Based on these preliminary results, we chose the lysostaphin lysibody as our lead molecule and LysK lysibody as a second option in further experiments.
Lysostaphin and LysK lysibodies bind a range of clinical S. aureus isolates and other Staphylococcus species.We used fluorescence microscopy to evaluate the binding of the lysostaphin and LysK binding domains to a range of clinically relevant S. aureus strains, including several methicillin- and vancomycin-resistant strains. To avoid nonspecific fluorescent signal resulting from binding of the lysibody Fc region to protein A on the surface of wild-type S. aureus cells, we fused each of the two binding domains to green fluorescent protein (GFP) in place of the Fc in this experiment. The two GFP fusion proteins showed cell wall-specific labeling of all S. aureus strains tested (see Fig. S1 in the supplemental material). Neither construct bound to the control organisms, Bacillus subtilis and Escherichia coli. Incubation with GFP alone did not result in fluorescent labeling of any of the strains.
For the lead molecule, the lysostaphin lysibody, we also used a modified competitive ELISA to measure its binding to a range of other bacterial species. In this assay, a set amount of the lysostaphin lysibody was preadsorbed onto various bacterial species in solution at optical density at 600 nm (OD600) values of 15, 10, 5, or 1 in phosphate-buffered saline (PBS). Following centrifugation of the bacteria, the amount of unbound lysostaphin lysibody in the supernatant was determined by a modified ELISA, as described above (see Fig. S2 in the supplemental material). This assay demonstrated that the lysostaphin lysibody bound with high affinity to S. aureus, Staphylococcus epidermidis, Staphylococcus hyicus, and Staphylococcus lugdunensis, and with somewhat lesser affinity to Staphylococcus sciuri. Little to no binding was observed for Micrococcus luteus, B. subtilis, Bacillus anthracis, Lactococcus lactis, S. pyogenes, Streptococcus agalactiae, Streptococcus bovis/gallolyticus, Enterococcus faecalis, Enterococcus faecium, and E. coli, demonstrating high specificity to the genus Staphylococcus.
Lysostaphin and LysK lysibodies promote robust deposition of complement on the surface of S. aureus.We next determined the ability of lysostaphin and LysK lysibodies to fix complement on the surface of S. aureus, using immunofluorescence microscopy. S. aureus cells were incubated with lysibodies or controls, then treated with human complement, and finally washed, fixed, and blocked. The extent of complement deposition on staphylococci was evaluated by fluorescence microscopy, using C3-specific antibodies and fluorescent conjugates. Lysostaphin and LysK lysibodies induced robust complement fixation on the surface of S. aureus, while controls PlyG lysibody (specific for B. anthracis), Fc alone, 1K8 (nonspecific monoclonal antibody), or PBS alone showed no activity (Fig. 4). No signal was observed with any of the lysibodies when complement was omitted.
Lysostaphin and LysK lysibodies fix complement on the surface of S. aureus. Complement deposition on the surface of S. aureus Wood 46 (protein A negative) was determined by fluorescence microscopy. Staphylococci were attached to cover slides and incubated with lysibodies and then with S. aureus-adsorbed human complement. The cells were then washed, fixed, and blocked. Complement was detected using specific antibodies and Alexa Fluor 594 conjugate; DNA was stained with 4′,6-diamidino-2-phenylindole (DAPI). Slides were imaged using deconvolution microscopy, and images are presented as maximum intensity projections.
Lysostaphin and LysK lysibodies induce efficient phagocytosis and killing of S. aureus.We expanded our preliminary macrophage phagocytosis analysis for the lysostaphin and LysK lysibodies and compared their activity to that of the ClyS lysibody, the most active lysibody from our previous study (31). The lysostaphin lysibody induced robust phagocytosis of S. aureus by Raw 264.7 macrophage, and it was effective at lower doses compared to the ClyS lysibody (Fig. 5A). A higher concentration of the LysK lysibody was required to achieve maximum phagocytic activity compared to the other two lysibodies. Similar results were obtained using peritoneal murine macrophages; the lysostaphin lysibody promoted efficient phagocytosis of S. aureus at substantially lower concentrations compared to the LysK lysibody (Fig. 5B). We also tested the ability of lysibodies to induce the killing of phagocytosed staphylococci (Fig. 5C). Following a 3-hour incubation with macrophages in suspension, lysostaphin and LysK lysibodies induced the killing of over 90% of staphylococci, in line with the previously characterized ClyS lysibody (31) (Fig. 5C). The PlyG lysibody and Fc-only controls did not lead to statistically significant killing of staphylococci.
Lysostaphin and LysK lysibodies induce the phagocytosis and killing of S. aureus by macrophages. Raw 264.7 macrophages (A) or peritoneal murine macrophages (B) were incubated with S. aureus Newman/pCN57 (GFP) in the presence of lysibodies at different concentrations. Percent phagocytosis was determined by flow cytometry. Experiments were done in duplicates; the error bars represent standard deviation. (C) Cells of S. aureus strain Newman were incubated with Raw 264.7 macrophages in suspension for 3 h in the presence of 10 μg lysibodies or controls. Killing compared to that in PBS control is presented. Experiments were preformed in triplicates, with three technical repeats for each biological repeat. Standard deviation values are presented and P values compared to the PlyG lysibody control were calculated using a t test; ** indicates P < 0.01.
We next examined the ability of lysostaphin and LysK lysibodies to promote the phagocytosis of various S. aureus strains by neutrophils (Fig. 6). When incubated with HL-60 neutrophils, lysostaphin and LysK lysibodies induced phagocytosis of S. aureus strains Wood 46 (methicillin-susceptible Staphylococcus aureus [MSSA], protein A negative), USA300 (MRSA), and USA600 (MRSA/vancomycin-intermediate Staphylococcus aureus [VISA]) in a complement-dependent manner, while controls had no activity (Fig. 6A). Similar results were obtained using human peripheral blood polymorphonuclear cells (PMNs); lysostaphin and LysK lysibodies induced phagocytosis of all staphylococcal strains tested in a complement-dependent manner, while controls had no effect (Fig. 6B). We next determined the dose-response curve for each lysibody. With all tested strains, the lysostaphin lysibody was able to induce phagocytosis of staphylococci at the lowest concentration, followed by the ClyS and LysK lysibodies (Fig. 6C). Nevertheless, for some strains, the LysK lysibody induced more efficient overall phagocytosis at high concentrations compared to the lysostaphin lysibody. Thus, the lysostaphin lysibody performed better than the ClyS lysibody in both macrophage and neutrophil phagocytosis assays, demonstrating the best in vitro results to date.
Lysostaphin and LysK lysibodies induce phagocytosis of S. aureus by neutrophils. Neutrophils were incubated for 1 h with FITC-labeled S. aureus strains Wood 46, USA300, and USA600, in the presence of lysibodies and 0.5% complement unless otherwise noted. Percent phagocytosis was determined by flow cytometry; experiments were done in triplicates. (A) Lysibodies induce the phagocytosis of S. aureus by HL-60 neutrophils in a complement-dependent manner; 5 μg lysibody were used per assay. Significance of P values was designated as follows: **, P < 0.01, and ***, P < 0.001. (B) Lysibodies induce the phagocytosis of S. aureus by human PMNs in a complement-dependent manner; 5 μg lysibody were used per assay. (C) Effect of lysibody dose on phagocytosis of S. aureus by HL-60 neutrophils.
Lysostaphin lysibody protects mice from challenge with MRSA.We next tested the ability of the lysostaphin lysibody to protect mice from challenge with MRSA/VISA strain USA600. Mice were injected intraperitoneally with 1 mg of the lysostaphin lysibody and 4 h later, were challenged intraperitoneally with 5 × 106 S. aureus USA600. Mouse viability was monitored daily for 4 days (Fig. 7A), at which time surviving mice were sacrificed and the bacterial load in the kidneys was evaluated (Fig. 7B). Mice treated with the lysibody had improved survival (91%) compared to control mice (42%). Furthermore, 8 of the 10 surviving lysostaphin lysibody-treated mice had no detectable bacteria in their kidneys, whereas all surviving control mice had bacteria in their kidneys, ranging between 105 to 109 CFU/g.
The lysostaphin lysibody protects mice from MRSA in a kidney abscess model. Five-week-old female BALB/c mice were injected with 1 mg lysostaphin lysibody (n = 11) or with PBS as control (n = 12). Four hours later, the mice were injected i.p. with 5 × 106 S. aureus strain USA600 (MRSA/VISA) cells. Mouse viability is presented in panel A; P value was calculated using log-rank. On the fourth day, surviving mice were sacrificed and the kidneys were removed and homogenized. Bacterial load was determined through serial dilution and plating, and the bacterial load per gram of kidney tissue is presented in panel B; P value was calculated using the Mann-Whitney test. Data are aggregated from four separate experiments.
Evaluation of the lysostaphin lysibody half-life in mice.Next, we determined the rate of lysibody clearance from mice. Four mice were each injected with 200 μg of the lysostaphin lysibody. Of these, two were injected intraperitoneally (i.p.) and two were injected intravenously (i.v.) through the tail vein. Lysibody concentration in the mouse serum was evaluated by ELISA. Both methods of injection resulted in a similar initial concentration in the blood, around 25 μg/ml 3 h following injection. Lysibody concentration then dropped to around 12 μg/ml after the first 48 h, and then declined after 120 h to 0.8 μg/ml and 3.1 μg/ml for the i.v. route, and 0.1 μg/ml and 1.61 μg/ml for the i.p. route (Fig. 8).
Half-life of the lysostaphin lysibody in mouse blood. FVB/NJ female mice were injected with 200 μg lysostaphin lysibody; two mice were injected intraperitoneally and two were injected intravenously. Blood was collected at 3 h, 48 h, and 120 h following injection, and antibody concentration in the serum was determined by capture ELISA.
Evaluation of preexisting antibodies against the lysostaphin binding domain in the human population.To evaluate the presence of preexisting antibodies against the binding domains of lysostaphin and LysK in humans, serum was obtained from 14 healthy volunteers. We also obtained two separate lots of commercially available pooled human serum for this purpose. Sera were reacted against Western blots containing GFP fusions to lysostaphin and LysK binding domains, purified by one-step metal-affinity chromatography. These blots also included GFP (purified in the same manner) as a negative control and tetanus toxin, for which most people have some level of antibodies due to vaccination, as a positive control (see Fig. S3 in the supplemental material). As expected, many of the sera from individual donors, as well as both serum pools, reacted with tetanus toxin (two bands with molecular weights of 50 kDa and 100 kDa are visible). None of the sera, however, reacted with lysostaphin and LysK binding domains. In the few cases where bands appeared in these lanes, they did not correspond to the correct molecular weight and likely represent immunity against an E. coli protein that was not entirely removed by single-step purification.
DISCUSSION
In this study, we created six lysibodies with binding domains from lysostaphin, LysK, PlySa4, PlySa6, PlySa7, and PlySa32, all of which bound to their target staphylococci. Of these, lysostaphin and LysK lysibodies were the most promising lead candidates. Both lysibodies bound to the S. aureus cell wall, induced complement fixation, and led to robust phagocytosis of S. aureus by macrophages and neutrophils. Lysostaphin lysibody had superior activity compared to the LysK lysibody as well as to the ClyS lysibody—the best lysibody described thus far. Lysostaphin lysibody was tested in a mouse model, demonstrating effective protection of mice from a MRSA/VISA challenge.
Vaccines and therapeutic antibodies are actively developed as an alternative to antibiotics in the treatment of MRSA infections (20, 21, 36–39). Despite concentrated efforts to create antibody-based solutions for MRSA in recent years, no vaccine has yet been approved for clinical use. StaphVax, a bivalent conjugate vaccine against the T5/T8 capsule types, showed no efficacy in a phase III clinical trial with a protection from bacteremia endpoint (40). Similarly, V710, a vaccine against iron surface determinant B (IsdB) did not reduce serious postoperative S. aureus infections in patients undergoing cardiothoracic surgery (19). Several monoclonal antibodies have also been tested in late-stage clinical trials; however, none of these studies has met its clinical endpoint (20).
The lack of clinical success of therapeutic candidates with favorable preclinical results is surprising and has led to the reexamination of the correlates of protection in humans (20). However, a possible explanation is that for a pathogen as complex as S. aureus, targeting a single surface protein may not be sufficient to drive effective protective immunity against the variety of strains encountered in the clinic or community (41). S. aureus is not only a highly evolved pathogen but is also a commensal of 30% of the human population (42). It possesses an impressive array of redundant virulence factors, and the specific subset of virulence factors often varies between different isolates (43). Proteomic experiments conducted on the surface proteins of S. aureus revealed great variability in the proteins expressed by different strains (44). Secreted proteins are similarly variable, showing a high degree of genome plasticity and an even more pronounced variability in secreted proteins detected in the medium (45). Moreover, both surface-attached and secreted virulence factors are under tight regulation in S. aureus, and their expression levels differ in different growth stages or in different niches in the infected host (46). Regulation of virulence factor expression may also provide an escape mechanism from therapeutic antibodies, as selective pressure due to exposure to an antibody could favor variants with lower expression level of the target; in-host adaptation has been documented in several cases for S. aureus (47–49). The S. aureus capsule is also not an ideal target, as not all strains possess a capsule, most notably USA300, the most prevalent MRSA strain in the US (50). These data highlight several drawbacks to the single-target approach for opsonic monoclonal antibodies against S. aureus. Some therapeutic antibodies now in clinical trials, such as MEDI4893 and ASN100, focus on neutralization of staphylococcal toxins and have shown promising results in certain infection settings, such as that of staphylococcal pneumonia, where such toxins are particularly important (51, 52). Nevertheless, development efforts toward the production of an effective opsonic vaccine continue, and they now focus on multivalent vaccines with several surface targets in order to achieve better efficacy and improved strain coverage (41). While lysibodies are essentially single-target monoclonal antibodies, their carbohydrate targets are highly conserved and abundant compared to the protein targets of most monoclonal antibodies and thus may enable improved strain coverage and a better outcome. The carbohydrate targets of lysibodies are conserved not only among different strains of S. aureus but also among coagulase-negative staphylococci. Moreover, surface carbohydrates play an essential role in cell wall structure and function (26); certain surface carbohydrates are essential for viability (53), while others play an important role in pathogenesis (54, 55). Escape mutants with altered carbohydrate structure are therefore less likely to occur compared to protein targets. Thus, a single lysibody could potentially be effective against a range of staphylococcal strains that far exceeds that achievable with most protein-directed monoclonal antibodies.
The lead lysibody candidate described in this work contains the binding domain of lysostaphin (33). The parental lysostaphin molecule targets S. aureus, and, to various degrees, also coagulase-negative staphylococci (56–58). Importantly, lysostaphin can kill both dividing and nondividing staphylococci (59). Lysostaphin was active against S. aureus strains resistant to methicillin (56, 60), as well as those resistant to vancomycin (61). Methicillin-resistant coagulase-negative staphylococci were similarly sensitive (57). Von Eiff et al. used the disc diffusion method on 429 S. aureus isolates, and all proved sensitive (60). Yang et al. similarly demonstrated that all 168 MSSA and 89 MRSA isolates tested were sensitive to lysostaphin (62). Thus, our observation that the lysostaphin lysibody binds various coagulase-positive and coagulase-negative staphylococcal isolates is consistent with a large body of published work on the parental lysostaphin enzyme.
Despite the susceptibility of S. aureus clinical isolates to lysostaphin, a few mutants with reduced susceptibility to lysostaphin were documented, including mutation of the femAB genes, the lyrA gene (63), and the sspC gene (64). Like phage lysins, the intact lysostaphin molecule contains separate domains responsible for cell wall binding and cleavage of the peptidoglycan. Resistance to lysostaphin could arise by inhibiting either of these functions; however, only resistance linked to the cell wall binding domain would also affect the lysostaphin lysibody binding. Of the mutations noted above, the femAB mutation could affect lysostaphin lysibody binding; however, this mutation drastically alters the cell wall structure and renders the organism susceptible to antibiotics, such as methicillin and oxacillin (65, 66). Mutation of the lyrA gene did not affect the binding of the lysostaphin binding domain to the cell wall and is therefore not likely to prevent lysostaphin lysibody binding. Not enough data are available to determine the effect of the sspC gene mutation on lysostaphin lysibody binding; however, this mutation led to severe pleotropic effects that would likely render the mutant nonpathogenic (64). Thus, while resistance is a possibility, it is evident that altering the binding target of the lysostaphin lysibody in the cell wall carries a high fitness cost, specially in the presence of beta-lactam antibiotics (66). Future work would address specific potential mechanisms of resistance and their implications for treatment of MRSA infection.
As with any biologic, there exists a potential for the development of antidrug antibodies following prolonged exposure (67). Previous studies have shown that repeated injection of the intact lysostaphin molecule into rabbits indeed resulted in the production of antibodies; however, these did not completely neutralize lysostaphin, as evident by detectable lytic activity remaining in the rabbit blood (56). Similarly, administration of lysostaphin was effective in the treatment of S. aureus infection in a rabbit endophthalmitis model, even following immunization of the rabbits with lysostaphin and the presence of an elevated anti-lysostaphin antibody titer (68). Nevertheless, observations made with the intact lysostaphin molecule only bear partial relevance for the lysostaphin lysibody, as this molecule only contains the lysostaphin cell wall binding domain. In fact, recent studies suggest that the catalytic domain of lysostaphin is the major driver of immunogenicity, while the binding domain is far less immunogenic (69). The molecule could be further deimmunized through elimination of T effector epitopes (70). Even if antibodies against the lysostaphin lysibody are produced, they are unlikely to reach substantial levels before the pathogen is cleared and would thus not hinder the course of treatment.
In conclusion, the lysibody approach represent a new and reproducible way to create effective immunity against conserved cell wall carbohydrates on the surface of Gram-positive bacterial pathogens, leading to their elimination. In this paper, we expanded the range of molecules that could be used to create lysibodies to also include bacteriocins, in addition to bacteriophage lysins and bacterial autolysins, further validating the concept and creating a lysibody with superior characteristics. Today, antibiotics are the only treatment option available on the market for staphylococcal infections, and staphylococci are becoming increasingly resistant to standard-of-care antibiotics (2). Lysibodies could be used to treat severe, life-threatening infections alone or in combination with drugs or as a prophylaxis to prevent such infections in presurgical patients. Thus, the lysibody approach represents a promising solution for staphylococcal infections, as well as for those caused by a range of other Gram-positive pathogens.
MATERIALS AND METHODS
Ethics statement.Samples from human subjects were obtained in accordance with a protocol approved by the Rockefeller University Institutional Review Board (institutional review board [IRB] number VFI-0790), and all subjects gave informed consent. Mouse work was performed in accordance with protocol number 14691H, approved by the Rockefeller University Institutional Animal Care and Use Committee (IACUC).
Fluorescence microscopy.Fluorescence microscopy was performed essentially as described previously (71). In brief, bacteria were fixed using 2.6% paraformaldehyde and 0.012% glutaraldehyde in growth medium containing 30 mM phosphate buffer (pH 7.4) for 15 min at room temperature and 30 min on ice. Bacteria were washed, attached to poly-l-lysine-coated cover glass, and blocked with 10% normal goat serum. Bacteria were incubated with lysibodies at 2 μg/ml in PBS containing 2% bovine serum albumin (BSA) and 1% gelatin, and then with fluorescent conjugates diluted 1:1,000, each for 1 h at room temperature. Slides were mounted in 50% glycerol and 0.1% p-phenylenediamine in PBS (pH 8).
Phase-contrast microscopy and fluorescence microscopy were performed using a Nikon Eclipse E400 microscope, equipped with a Nikon 100×/1.25 oil immersion lens, and a Retiga EXi fast 1394 camera (QImaging). Image capture was done using QCapture Pro version 5.1.1.14 software (QImaging). Deconvolution microscopy was done on a DeltaVision image restoration microscope (Applied Precision/Olympus) equipped with CoolSnap quantum efficiency (QE) cooled charge-coupled device (CCD) camera (Photometrics). An Olympus 100×/1.40 numerical aperture (NA), UPLSApo oil immersion objective was used with a 1.5× optovar. Z-stack intervals were 0.15 μm. SoftWoRx software (Applied Precision/DeltaVision) was used for image deconvolution, and resulting images were corrected for chromatic aberrations. Experiments were repeated twice in duplicates.
Phagocytosis assays.Phagocytosis assays were performed essentially as described previously (31). In brief, 24-well plates containing confluent Raw 264.7 macrophages or peritoneal macrophages, derived from BALB/c female mice, were supplemented with 1 × 107 S. aureus Newman/pCN57 (expressing GFP) cells, and incubated for 1 h at 37°C, 5% CO2, at various lysibody concentrations. The wells were washed several times to remove extracellular bacteria, fixed with 1 ml of 1% paraformaldehyde in PBS for 1 h at 4°C, and washed. Raw 264.7 macrophages were scraped off the plate using a disposable loop in 200 μl of PBS, while peritoneal macrophages were incubated with 250 μl of 0.25% trypsin in PBS (pH 7.2) containing 0.1% EDTA for 30 min at 37°C, gently suspended by pipetting with a 1-ml pipette tip, and washed.
For killing experiments, each assay contained 105 log-phase S. aureus strain Newman cells and 10 μg lysibody in a total volume of 100 μl Hanks' balanced salt solution (HBSS) containing 0.1% gelatin, in a well of a 96-well U-bottom plate. The plate was placed on a shaker for 1 h at 200 rpm at 4C°. Raw 264.7 macrophages (105) were then added in a total volume of 100 μl HBSS (200 μl final volume), and the plate was placed on a shaker for 3 h at 200 rpm at 37C°. Samples were lysed in 0.2% saponin, serially diluted in distilled water, and plated for CFU quantification. Experiments were performed in triplicates and included three technical repeats for each biological repeat.
For phagocytosis assays with neutrophils, HL-60 neutrophils were prepared as described by Romero-Steiner et al. (72), and human peripheral blood PMNs were isolated from healthy volunteers as previously described (31). Neutrophils were suspended in Hanks' balanced salt solution (Gibco 14025) containing 0.1% gelatin at 1 × 108 cells/ml. Fluorescein isothiocyanate (FITC)-labeled bacteria were suspended to a final concentration of 5 × 107 cells/ml in HBSS containing 0.1% gelatin. Bacteria (10 μl) were added to each well of a U-bottomed 96-well plate containing 70 μl HBSS containing 0.1% gelatin and lysibodies. Following 1 h of incubation with shaking at 4°C, 10 μl of 5% S. aureus-adsorbed complement and 10 μl neutrophils at 1 × 108 cells/ml were added, and the plate was shaken at 200 rpm at 37°C for 1 h. The cells were then fixed with 2.6% paraformaldehyde for 1 h, blocked, and analyzed by flow cytometry, first by gating on neutrophils using forward and side scatter, and then by determining the percentage of neutrophils containing fluorescent S. aureus. Experiments were performed in triplicates and repeated three times.
Complement fixation.Complement fixation procedures were performed essentially as described previously (31). Briefly, staphylococci were attached to poly-l-lysine-coated cover slides and incubated with lysibodies at a final concentration of 1 mg/ml in Dulbecco's phosphate-buffered saline (DPBS) for 1 h at room temperature. The cells were washed with PBS and DGHB buffer (5 mM HEPES, 71 mM NaCl, 0.15 mM CaCl2, 0.5 mM MgCl2, 2.5% glucose, and 0.1% gelatin [pH 7.4]) (73) and then incubated for 20 min at 37°C with 30 μl DGHB containing 0.5% S. aureus-adsorbed human serum. The cells were washed with PBS and fixed with 2.6% paraformaldehyde in PBS for 1 h at 4°C. The cells were washed with PBS and blocked with PBS containing 2% BSA and 1% gelatin. C3b deposition was detected with rabbit anti-C3 diluted 1:500, followed by goat anti-rabbit Alexa Fluor 594 conjugate diluted 1:1,000 and 1 μg/ml 4′,6-diamidino-2-phenylindole (DAPI). Experiments were repeated twice in duplicates.
ELISAs.Plate preparation was performed as follows. High binding polystyrene 96-well plates were coated with 100 μl of 0.01% poly-l-lysine at room temperature for 1 h and washed with water. S. aureus strain Wood 46 (protein A negative) was grown overnight, washed, and resuspended in PBS to a final OD600 of 1.0. To each well of the microtiter plate were added 50 μl cell suspension and 50 μl freshly prepared 3.2% paraformaldehyde solution in PBS. The plates were incubated at room temperature for 20 min and centrifuged for 20 min at 1,500 × g at 4°C. The plates were washed with water three times and incubated with 150 μl/well 0.1 M lysine in PBS for 1 h at room temperature. The plates were then washed three times with water and twice with ELISA wash buffer (10 mM sodium phosphate, 150 mM NaCl, 0.05% Brij-35, and 0.02% sodium azide). The plates were then blocked with PBS containing 1% BSA overnight at 4°C.
For ELISA to compare the binding of different lysibodies to S. aureus, various lysibodies were serially diluted 2-fold from an initial stock of 10 μg/ml, and 50 μl of each dilution was transferred to a well of the prepared ELISA plate and incubated overnight at 4°C. The wells were then washed three times with water and twice with wash buffer and incubated with 100 μl goat anti-human IgG Fc alkaline phosphatase conjugate at 1:10,000 dilution for 3 h at 37°C. The plate was washed as described above, developed with p-nitrophenyl phosphate (pNPP), and absorption at 405 nm was measured using a SpectraMax Plus plate reader (Molecular Devices).
Determination of lysibody concentrations in mouse serum was done in a similar manner. Mouse serum was serially diluted in PBS containing 1% BSA and all dilutions were applied to an ELISA plate. Three serum dilutions resulting in values within the assay's dynamic range were used to calculate the serum lysibody concentration based on a lysostaphin lysibody standard found on the same plate.
For studies involving lysibody adsorption onto bacterial strains, each of the strains was grown overnight, washed, and suspended to a final calculated OD600 of 15, from which it was subdiluted to OD600s of 10, 5, and 1. In a U-bottomed 96-well plate that was preblocked with PBS containing 1% BSA, 50 μl of lysibody at 10 μg/ml were mixed with 50 μl of bacterial suspensions and incubated for 1 h at 37°C with shaking at 200 rpm. The plate was then centrifuged for 10 min at 4,000 rpm, and 75 μl of the supernatant was transferred to an ELISA plate. Preparation and processing of the ELISA plate was performed as described above. ELISAs were repeated at least twice.
Mouse kidney abscess model.The mouse kidney abscess model was modified from Raz et al. (31). Five-week-old BALB/c female mice (Charles River Laboratories) were injected i.p. with 1 mg lysibody or control. Four hours later, 5 × 106 CFU of S. aureus strain USA600 (MRSA/VISA) in saline containing 5% hog gastric mucin (Sigma) were injected to each mouse. Bacteria were prepared as follows: an overnight culture of USA600 was diluted 1:100 in brain heart infusion (BHI) and grown to an OD600 of 0.5 at 37°C with shaking at 200 rpm. Bacteria were washed in saline, suspended in saline to an OD600 of 1.0, and diluted to the desired concentration; actual CFU/ml were evaluated by plating. Mice were monitored daily, and after 4 days surviving mice were sacrificed. The kidney bacterial load was determined by aseptically removing the kidneys, grinding the kidneys in 1 ml of 0.5% saponin, performing serial dilutions, and plating.
Determination of lysibody half-life in mice.Four 3-month-old FVB/NJ female mice were each injected with 200 μg of lysostaphin lysibody. Two mice were injected intraperitoneally and two were injected i.v. through the tail vein. Blood was obtained by retroorbital bleeding at the 3-h, 48-h, and 120-h time points; serum was obtained by centrifugation following 1 h of coagulation. Quantification of the lysostaphin lysibody concentration present in mouse serum was done by ELISA as described above, using serially diluted purified lysostaphin lysibody as the standard.
Detection of preexisting immunity against lysostaphin and LysK binding domains in human serum.Blots for detection of antibody response in human serum were prepared by separating 250 ng of each protein on 10% SDS-PAGE and transferring the resulting gel to a polyvinylidene difluoride membrane. Blots were blocked with PBS supplemented with 5% nonfat dry milk and 0.1% Tween 20 (PBS-DT), washed with Tris-buffered saline, and incubated overnight with 10 ml PBS-DT containing 100 μl human serum. Blots were washed 3 times and incubated with goat anti-human Fc horseradish peroxidase conjugate diluted 1:5,000 in PBS-DT. Blots were developed using SuperSignal West Pico chemiluminescent substrate (Pierce).
Statistical analysis.A two-tailed Student's t test was used to evaluate statistical significance in phagocytosis assays. A log rank test was used to evaluate mouse survival data. Statistical significance for bacterial load in the kidneys was evaluated using the two-tailed Mann-Whitney test; the maximal bacterial load value observed in these experiments was assigned to mice that succumbed to infection. Prism version 5.0c (GraphPad Software, La Jolla, CA) was used for data analysis.
ACKNOWLEDGMENTS
We thank Yi Wu, Tiffany Benjamin, and Anaise Hernandez for help with various aspects of this work. We thank Svetlana Mazel and the members of the Rockefeller University Flow Cytometry Resource Center for their advice on flow cytometry. We are grateful to Alison North and members of the Rockefeller University Bio-Imaging Resource Center for their advice regarding fluorescence microscopy. We thank Ray Schuch for his thoughtful advice during the course of this study.
Author contributions: A.R. and V.A.F. designed the research; A.R., A.S., M.T., and T.A. performed the research; A.R., A.S., M.T., T.A., and V.A.F. analyzed data; and A.R. and V.A.F. wrote the paper.
We declare no conflict of interest.
FOOTNOTES
- Received 21 May 2018.
- Returned for modification 13 June 2018.
- Accepted 16 July 2018.
- Accepted manuscript posted online 23 July 2018.
Supplemental material for this article may be found at https://doi.org/10.1128/AAC.01056-18.
REFERENCES
- Copyright © 2018 American Society for Microbiology.
