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J. Biol. Chem., Vol. 281, Issue 8, 4831-4843, February 24, 2006

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Immunogenicity and Protective Efficacy of Bacillus anthracis Poly--D-glutamic Acid Capsule Covalently Coupled to a Protein Carrier Using a Novel Triazine-based Conjugation Strategy^*

Joseph Joyce¹, James Cook, Donald Chabot, Robert Hepler, Wesley Shoop^¶, Qiuwei Xu, Thomas Stambaugh^||, Miguel Aste-Amezaga, Su Wang, Lani Indrawati, Mark Bruner, Arthur Friedlander, Paul Keller, and Michael Caulfield

From the Departments of Vaccine and Biologics Research, Laboratory of Science and Investigative Toxicology and Bioprocess and Bioanalytical Research, Merck Research Laboratories, West Point, Pennsylvania 19486, United States Army Medical Research Institute of Infectious Diseases, Frederick, Maryland 21701, the ^¶Department of Human and Animal Infectious Disease Research, Merck Research Laboratories, Rahway, New Jersey 07065, and ^||Lockheed Martin, Edison, New Jersey 08837

Received for publication, August 25, 2005 , and in revised form, October 20, 2005.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The capsular polypeptide of Bacillus anthracis is composed of a unique polyglutamic acid polymer in which D-glutamate monomers are joined by -peptidyl bonds. The capsule is poorly immunogenic, and efforts at exploiting the polymer for vaccine development have focused on increasing its inherent immunogenicity through chemical coupling to immune-stimulating protein carriers. The usual strategy has employed carbodiimide-based condensing reagents for activation of free -carboxyl groups, despite reports that this chemistry may lead to chain scission. We have purified the high molecular mass capsule to >95% homogeneity and have demonstrated that the polymer contains >99% poly--D-glutamic acid. The predominant structure of the polymer as assessed by circular dichroism and multiangle laser light scattering was unordered at near-neutral pH. We investigated the effects of various activation chemistries, and we demonstrated that carbodiimide treatment under aqueous conditions results in significant cleavage of the -peptidyl bond, whereas scission is significantly reduced in nonaqueous polar solvents, although undesired side chain modification was still observed. An activation chemistry was developed using the triazine-based reagent 4-(4,6-dimethoxy (1,3,5)triazin-2-yl)-4-methylmorpholinium chloride, which allowed for controlled and reproducible derivatization of -carbonyls. In a two-pot reaction scheme, activated capsule was derivatized with a sulfhydryl-reactive heterobifunctional moiety and was subsequently coupled to thiolated carrier protein. This conjugate elicited very high capsule-specific immune titers in mice. More importantly, mice immunized with conjugated capsule exhibited good protection against lethal challenge from a virulent B. anthracis strain in two models of infection. We also showed, for the first time, that treatment of capsule with carbodiimide significantly reduced recognition by capsule-specific antisera concurrent with the reagent-induced reduction of polymer mass. The data suggested that for vaccine development, maintenance of the high mass of the polymer may be important.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Anthrax is a highly infectious disease caused by the Gram-positive spore-forming organism Bacillus anthracis (1). In humans, the infection can be lethal and typically occurs through one of three routes: cutaneous, gastrointestinal, or inhalation. Cutaneous anthrax is typically contracted through open skin wounds in contact with infected animal materials such as hides, wool, etc. It is the most common and least severe form of the disease, accounting for 95% of reported cases. Gastrointestinal infection is usually the result of ingestion of undercooked infected meat, and respiratory infection involves the inhalation of air-borne spores. High mortality rates are common with the latter two forms. Early symptoms of inhalational anthrax are nonspecific, but death can follow rapidly with little opportunity for therapeutic intervention (2).

Anthrax has long been considered one of the most likely bio-warfare weapons. Its potential use as an agent of bio-terrorism was dramatically highlighted in 2001 by a series of postal mailing attacks in the United States that resulted in 11 cases of cutaneous and 11 cases of inhalational anthrax, including 5 fatalities as a result of the latter (3). The most effective defensive strategy in the face of such an event is the availability of a highly efficacious and well tolerated vaccine.

Two major plasmid-derived bacterial virulence factors have been identified in B. anthracis. The first is the tripartite toxin encoded by plasmid pXO1 that is composed of lethal factor, edema factor, and their associated receptor-binding component, protective antigen (PA)² (4). The second is a surface capsular polypeptide composed of poly--D-glutamic acid (PGGA), the product of plasmid pXO2 (5). Historical vaccines such as the Pasteur and Sterne formulations were based on live attenuated strains, but the currently licensed vaccine, Anthrax Vaccine Adsorbed, is a sterile cell-free filtrate of bacterial culture medium adsorbed to an aluminum-based adjuvant. This vaccine has been shown to confer good protection in humans (6), but it is a fairly undefined formulation and requires a 6-injection dosing regimen. Furthermore, there has been an ongoing debate about its safety, although large trials of the vaccine have indicated an acceptable safety level (7, 8). As a result of these considerations, several approaches to a second generation vaccine are in development, the majority of which target PA. Such vaccines would provide indirect protection because they would act to neutralize the toxin following the establishment of infection rather than offering sterilizing immunity. A second bacterial component of interest is the capsule. This polypeptide is an attractive vaccine candidate for several reasons. First, acapsular strains of B. anthracis have been shown to be highly attenuated in a variety of animal models (9, 10). Second, the -peptidyl linkage and D-amino acid composition of the polymer are unique against the background protein repertoire of the host and would be expected to generate a highly specific immunological response. Finally, recent studies have demonstrated that antibodies against the capsule are opsonophagocytic, suggesting that vaccination with the capsule may produce a response that would target the bacillus directly and prevent establishment of initial infection, generating sterilizing immunity (11). This hypothesis is further supported by the demonstration of good passive protection in mice administered a capsule-specific monoclonal antibody and challenged in a lethal pulmonary model of infection (12).

Early studies indicated that PGGA isolated from B. anthracis is poorly immunogenic and confers little or no protective efficacy in animal models of infection (13). More recent work has confirmed that the capsule functions as a T-independent antigen (14), although the report by Chabot et al. indicated some protection in mice from a subcutaneous spore challenge with a pXO1^-, pXO2⁺ derivative of the Ames strain and enhanced protection against fully virulent pXO1⁺, pXO2⁺ Ames strain when given with PA (15). An immune-enhancing strategy, which has proved successful for related T-independent bacterial polysaccharide antigens, involves covalently coupling the biopolymer to a highly immunogenic protein carrier. These immunoconjugates often produce specific serum antibody titers several orders of magnitude higher than the antigen alone, and immune responses can often be manipulated depending on the choice of carrier or adjuvant (16). Recent literature reports have described this approach for PGGA where the carriers used were bovine serum albumin (BSA) (15), Pseudomonas aeruginosa exotoxin A (17), keyhole limpet hemocyanin (11), and PA (18). Mice immunized with these conjugates produced anti-capsule-specific sera that induced killing of toxin-minus, capsule-positive bacilli in an in vitro opsonophagocytic assay (17); furthermore, mice immunized with PGGA-PA conjugates were protected against lethal challenge by co-administered protective antigen and lethal factor toxin components (18).

Here we describe preparation of highly purified and well characterized conjugates of PGGA covalently coupled to the outer membrane protein complex (OMPC) of Neisseria meningitidis serotype B (19). We chose this carrier based on the fact that it is a component of a currently licensed pediatric vaccine against Haemophilus influenzae type B (20) and has been used as a carrier in investigational vaccines for other bacterial capsular polysaccharides, including several serotypes of Streptococcus pneumoniae (21). Furthermore, OMPC presents certain unique immune-enhancing effects that render it superior to similar protein-based carriers (22). It was our desire to produce a highly purified PGGA as starting material and to avoid excessive antigen processing during purification and conjugation in the event that conformational epitopes present on high molecular weight material might be important neutralizing determinants. The purification and conjugation protocols described herein met these goals. We developed a coupling strategy that allowed for precise and controllable derivatization of PGGA and OMPC separately, an important consideration in being able to exercise fine control over modification of the antigen and achievable loading levels of polypeptide on carrier. Finally, we tested our conjugates in two mouse challenge models, utilizing vegetative and spore forms of B. anthracis, and we evaluated the passive protective efficacy of the polyclonal animal serum.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Bacterial Strain, Inoculation, and Culture Methods—B. anthracis strain Ames (United States Army Medical Research Institute of Infectious Diseases) was streaked for isolation onto Miller LB (Difco; 10 g of pancreatic casein digest, 5 g of yeast, 10 g of sodium chloride, 15 g of agar) agar plates from a glycerol freeze-down and grown overnight at 37 °C. A single bacterial colony was used to inoculate 5 ml of LB broth in a 50-ml polyethylene disposable centrifuge tube. The culture was incubated overnight in an air shaker at 37 °C. NBY/bicarbonate plates were prepared from 8g of nutrient broth powder (Difco), 12 g of agar, and 3 g of yeast extract in 900 ml of water. The medium was autoclaved, and 100 ml of an 8% 0.45-µm filtered sodium bicarbonate solution was then added prior to plates being poured. A total of 20 NBY/bicarbonate plates were streaked using an inoculating loop with the overnight Ames culture, and the plates were incubated at 37 °C with 20% CO₂ for 24–48 h.

Isolation of Crude PGGA—Bacteria were removed from confluent plates using a disposable spreader and resuspended in water. The suspension was autoclaved at 121 °C for 1 h and clarified by centrifugation at 11,000 x g for 10 min at 4 °C. The supernatant liquid was recovered, and sterility was verified. The extract was buffered by addition of 0.1 volume of 0.1 M Tris, pH 8.0, 0.1 M sodium chloride, 0.01 M magnesium chloride, 0.2% (w/v) sodium azide. Nuclease digestion was performed by addition of 100 units of DNase I (Invitrogen) and 10 units of RNase ONE (Promega, Madison, WI) overnight at 4 °C. Trichloroacetic acid (50% w/v) was added to a final concentration of 5%, and the mixture was incubated on ice for 30 min. The preparation was clarified by centrifugation at 12,000 x g for 15 min at 4 °C, and the supernatant was recovered. The pH was adjusted to 7 by slow addition of 6 N NaOH, and the material was exhaustively dialyzed against water for 3 days at 4 °C. The crude product was lyophilized to dryness, and a second sterility confirmation was performed.

Polishing of Crude PGGA—Crude PGGA was dissolved in water at 2 mg/ml, and the solution was mixed with an equal volume of hydroxyapatite (HPT) chromatography Buffer A (0.004 M sodium phosphate, pH 7.0, 1.0 M NaCl). Sample was applied to a column (25 mm inner diameter x 200 mm) of HPT type II (Bio-Rad) at a flow rate of 8 ml/min on a Waters Delta 600 (Milford, MA) chromatography system. Elution was monitored by absorbance at 215 nm. Nonbound material was washed out with Buffer A, and PGGA was eluted with a linear gradient from 0 to 100% Buffer B (0.4 M sodium phosphate, pH 7.0, 1.0 M NaCl) at 5 ml/min. Fractions (8 ml) were collected during elution, and those containing PGGA by analytical size exclusion chromatography were pooled. The HPT product was concentrated and diafiltered against water using a 5-kDa PES Pellicon XL50 tangential-flow ultrafiltration membrane (Millipore, Billerica, MA) to give the final polished product. Purified PGGA was shell-frozen, lyophilized to dryness, and stored desiccated at -70 °C.

NMR Analysis—All analyses were performed on a 600-MHz Varian NMR instrument. Crude or purified PGGA powder was weighed and dissolved in a fixed volume of D₂O (99.999%) containing 0.01% Me₂SO of known concentration for the quantification of polypeptide concentration and purity. The spectral chemical shift was internally referenced with 0.02% d₆-2,2-dimethyl-2-silapentanesulfonic acid. The acquisition was carried out in 5-mm tubes at a probe temperature of 25 °C. NMR diffusion spectra were generated with the pulse sequence, Bi-polar Longitudinal Eddy Current Delay (23), at a pulsed field gradient of 0.18 torr/m and 2-ms duration. Two-dimensional HMBC NMR of PGGA was recorded at 25 °C. The spectral width of the proton and carbon dimensions were 4001.6 and 15080.1 Hz, respectively. Because of the limited spectral width, the carbon chemical shift of carbonyl carbon was folded into the spectrum. The size of the acquired spectral matrix was 2416 x 128, and the 90° pulse of proton and carbon was 7 and 17 µs, respectively. The delay for the development of one and multiple bound proton-carbon correlation was optimized for 145 and 10 Hz, respectively. The raw data were multiplied with 90° shifted sine wave function along both dimensions. The final spectrum matrix after Fourier transformation was 2048 x 2048.

Determination of Absolute Configuration—Purified PGGA (10 µl) was dried under vacuum in a sealed reaction vessel for 20 min using a Pico-Tag Work station (Waters). The residue was solubilized in 400 µl of 6 N constant boiling HCl and incubated under nitrogen at 110 °C for 20 h. Following hydrolysis, 200 µl of HPLC-grade water was added; the reaction tube was vortexed, and the mixture was transferred to a clean vial. The wash step was repeated four times, and the diluted hydrolysate was dried in a speed-vac concentrator. The residue was taken up in 200 µl of (R)-2-butanoic HCl (5:1 v/v mixture of R)-2-butanol (Sigma) and acetyl chloride) and incubated at 120 °C for 20 min. Residual (R)-butanoic HCl was evaporated under nitrogen. Heptafluorobutyric anhydride, 100 µl, was added, and the mixture was incubated at 120 °C for 10 min. The solvent was removed under nitrogen, and the N-heptafluorobutyryl (R)-2-butyl diester derivative of glutamic acid was recovered in a mixture of 100 µl of ethyl acetate and 100 µl of acetic anhydride before being transferred to a gas chromatography (GC) autosampler vial. As controls, D- and L-glutamic acid (Sigma) were prepared at 0.7 µg/ml in methanol, and a 25-µl aliquot of each was derivatized as for PGGA. GC analysis was performed on a Hewlett-Packard HP6890 with flame ionization detector (FID). The column was an HP-5MS (30 x 0.32 mm) coated with 5% phenyl, 95% dimethylpolysiloxane (Agilent, Palo Alto, CA). Temperatures of injection port and FID were maintained at 250 and 300 °C, respectively. The temperature of the GC oven was programmed starting from 80 °C, holding for 5 min, ramping at a rate of 8 °C/min to 275 °C, and holding for 2 min. Helium gas, at a flow rate of 2.2 ml/min, was utilized as the mobile phase. Sample injection volume was 2 µl.

Sizing Analyses—Molecular mass determination of PGGA was performed by high performance size exclusion chromatography (HPSEC) coupled with multiangle laser light scattering (MALLS) detection on an 1100 series liquid chromatography system (Agilent) and Dawn^TM EOS 18 angle light scattering detector with quasi-elastic light scattering and Optilab^TM DSP options (Wyatt, Santa Barbara, CA). Chromatography was performed at a flow rate of 0.5 ml/min using two Ultrahydrogel^TM Linear 30-cm columns in series behind an Ultrahydrogel^TM guard column (Waters). Running buffer was 50 mM sodium phosphate, pH 7.2, 0.15 M sodium chloride containing 8 ppm Proclin 150^TM as a preservative. Columns and detectors were maintained at 35 °C. The refractive index increment (dn/dc) of purified PGGA was determined empirically. Briefly, five PGGA solutions of approximate concentration 0.2–0.8 mg/ml were prepared in phosphate-buffered saline and dialyzed exhaustively against the same buffer. The refractive index of each sample (0.2 µm filtered) was measured using a Optilab DSP interferometric refractometer (Wyatt), and concentration was determined from the absorption at 215 nm using an empirically determined extinction coefficient of 7628. The concentration was input into the Dn/Dc software (Wyatt Technology DNDC version 5.90.03) to obtain a value of 0.150 ± 0.002, which was used for all subsequent mass determinations. PGGA sonication studies were performed in 50 mM phosphate, 0.15 M NaCl, pH 7.5, using a Branson Ultrasonics Sonifier (St. Louis MO) with either 2-inch or high intensity cup/horn probes. Sonication was carried out at an output energy of 180 watts.

Amino Acid Analysis—Quantitative amino acid analysis was performed using the AccQTag^TM system (Waters). PGGA samples were appropriately diluted in water and hydrolyzed in constant boiling 6 N HCl containing 2% phenol at 110 °C for either 20 or 70 h, dried, and reconstituted in 20 mM HCl. Hydrolysates were labeled with AccQFlu- or^TM reagent following the manufacturer's protocol and heated at 55 °C for 10 min prior to analysis. Chromatography was performed on an 1100 series HPLC (Agilent) using an AccQTag^TM Amino Acid Analysis Column at a flow rate of 1.0 ml/min. Fluorescence was monitored at excitation and emission wavelengths of 250 and 395 nm, respectively.

CD Analysis—CD measurements were performed on a J-810 spectropolarimeter equipped with a Peltier temperature controller set at 25 °C (Jasco, Easton, MD). Spectra were collected from 185 to 300 nm at a scan speed of 100 nm/min for 10 accumulations/sample with a data interval of 0.1. The polymer concentration of samples was accurately quantified by amino acid analysis, and the CD signal was converted to molar ellipticity, [], as shown in Equation 1,

(Eq. 1)

where MRW is the mean residue weight, which in this case was the polymer repeat unit molecular mass (128.1Da); l is path length in cm, and c is concentration in mg/ml.

Nucleic Acid Analysis—PGGA samples were dissolved in water at a concentration of 1 mg/ml (w/v) and analyzed for total nucleic acid content using the PicoGreen^TM fluorescent dye-binding assay (Molecular Probes, Eugene, OR) as per the manufacturer's recommendations. Quantitation was performed by using either calf thymus DNA or total yeast RNA as nucleic acid standards.

Aqueous Carbodiimide Activation of PGGA—Aqueous activation studies were performed by dissolving PGGA to a final concentration of 4 mg/ml in either 0.25 M sodium phosphate, pH 7.3, or 0.25 M MES, pH 4.5. Both buffers contained 0.15 M NaCl and 0.75 mg/ml N-(-maleimidocaproic acid)hydrazide (EMCH, Pierce). 1-Ethyl-3(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC, Pierce) was added to final concentrations of 5 or 0.5 mg/ml. Reactions were allowed to proceed at room temperature for 1–3 h in the dark. At each time point, the reactions were stopped by three consecutive rounds of buffer exchange over a 5,000-Da centrifugal concentrator Samples were analyzed by HPSEC/MALLS, and aliquots were removed for ELISA testing, snap-frozen in liquid nitrogen, and stored at -70 °C. Untreated controls were prepared by dissolving PGGA in either 0.25 M sodium phosphate, pH 7.0, 0.15 M sodium chloride, or 0.25 M MES, pH 4.5, 0.15 M sodium chloride. The controls were analyzed at 0 and 3 h.

Development of Nonaqueous Activation Chemistry—In order to conserve purified PGGA, initial studies were performed using poly--D-glutamic acid (PAGA) (Sigma) and subsequently confirmed with PGGA. In order to render the polymers soluble in nonaqueous solvents, they were first converted from the sodium to tetrabutylammonium (TBA⁺) salt by chromatography on AG50W-X8 resin (Bio-Rad). Resin (200–400 mesh, H⁺ form) was washed in distilled deionized water (DI water) and used to pack a 10-ml column that was washed successively with 10 volumes water, 1 M tetrabutylammonium hydroxide, and water. PAGA or PGGA dissolved in DI water (3–5 mg/ml) was applied at 1 ml/min and eluted with DI water. The eluent was dried and product yield determined by weight. The TBA⁺ adduct was reconstituted in anhydrous dimethylformamide (DMF) at a nominal concentration of 10 mg/ml and placed on ice. EMCH was added at a equimolar ratio to total carboxyl content. For comparison of condensing reagents, either N,N'-diisopropylcarbodiimide (DIPC) or 4-(4,6-dimethoxy(1,3,5)triazin-2-yl)-4-methylmorpholinium chloride (DMTMM; Acros Organics, Pittsburgh, PA) dissolved in DMF was added at an equimolar ratio to carboxyl. For the DIPC reaction, 1-hydroxybenzotriazole was added at a 0.1 molar eq to carboxyl. Reactions proceeded in dark under N₂ for 1 h on ice followed by ambient temperature overnight. Activated PAGA was desalted by gel permeation chromatography in DMF using Sephadex® LH-20 resin (Amersham Biosciences), and the product was diluted with 4 volumes DI water. The product was dialyzed successively versus 1 M NaCl and DI water and dried under vacuum. For comparison of maleimidation reagents, either EMCH or 5-aminopentylmaleimide (APM) was added to PGGA dissolved in DMF at varying molar ratios (0.2 to 1) to carboxyl. DMTMM (0.2–1 eq) was added, and reactions were carried out under N₂ in dark on ice for 1 h followed by ambient temperature for 3 h. Product was counter-ion exchanged by dialysis as described above, omitting the gel permeation step, and dried under vacuum. For evaluation of DMTMM under aqueous conditions, PGGA was dissolved in 0.1 M HEPES, pH 7.3, 0.5 M NaCl, 2 mM EDTA buffer at 3 mg/ml, with addition of EMCH or APM and subsequent reaction conditions as described above.

Preparation of PGGA-OMPC Conjugate—Purified outer membrane protein complex (OMPC) of N. meningitidis serotype B was activated by thiolation of lysine residues as described previously (24). Total sulfhydryl content of the activated carrier was determined using 5,5'-dithionitrobenzoic acid (25). PGGA maleimidation was optimized based on activation studies. Briefly, PGGA (TBA⁺) was dissolved in DMF at 5 mg/ml, and EMCH was added at an equimolar equivalent to carboxyl. After 10 min, DMTMM was added at a 0.2 molar eq, and the reaction was purged with N₂ and incubated on ice in the dark for 1 h followed by 2.5 h at ambient temperature. The reaction mixture was diluted 4-fold with DI water and dialyzed as described above in the dark at 4 °C, with continuous N₂ purging of the dialysis buffer. The maleimide-adducted product was 0.22-µm filtered and buffered to a final concentration of 20 mM HEPES, pH 7.3, 0.5 M NaCl, 2 mM EDTA. Thiolated OMPC was added at an equimolar ratio of sulfhydryl to maleimide, and the reaction was allowed to proceed for 28 h at ambient temperature in the dark. An OMPC-only control was carried forward in parallel. Residual unreacted thiol and maleimide groups were successively quenched using a 5-fold molar excess of iodoacetamide and N-acetylcysteamine, respectively, overnight at ambient temperature in the dark. To remove excess reagents and residual free PGGA, products were pelleted by centrifugation at 289,000 x g for 60 min at 4 °C and resuspended in 20 mM HEPES, pH 7.3, 0.15 M NaCl buffer. This step was repeated twice, and the final resuspension was in 0.15 M NaCl. Product and control were centrifuged at 1,000 x g for 10 min to remove any aggregated material. Protein was determined by modified Lowry assay (26). Conjugate and control were adsorbed onto aluminum hydroxyphosphate adjuvant (alum, Merck) and diluted to appropriate PGGA concentrations in sterile saline.

Animal Immunizations—BALB/c mice were immunized by intraperitoneal injection of 0.2 ml of alum-formulated conjugate vaccines.

PGGA ELISA—Serum IgG antibody titers against PGGA were measured by ELISA as follows. Costar high binding plates were coated with 2 µg/ml (50 µl/well) purified PGGA in phosphate-buffered saline and incubated overnight at 4 °C. Plates were blocked overnight at 4 °C with 5% fetal bovine serum. Serum samples were tested at 1:5 serial dilutions on the ELISA plates after they were prediluted at 1:10 or 1:100. The plates were then incubated overnight at 4 °C. After washing, alkaline phosphatase-labeled goat anti-mouse IgG (Southern Biotech, Birmingham, AL) at 1:2,000 dilution was used to detect bound IgG antibody. The plates were developed using p-nitrophenyl phosphate substrate (Sigma), and absorbance was measured at 405 nm. Titers were calculated as described previously (27).

Animal Challenge Studies—BALB/c mice, immunized as described above, were challenged with either B. anthracis vegetative or spore preparations via the intraperitoneal route. B. anthracis Ames strain spores were maintained at -70 °C. To prepare the vegetative bacillus inoculum, the frozen spore stock material was thawed, and a loop of material was streaked on a sheep red blood agar plate and incubated for 18 h at 35 °C. Immediately prior to challenge, the 18-h vegetative bacillus colonies were suspended in saline, vortexed vigorously, and diluted to achieve an inoculum shown previously to provide a specific number of colony-forming units (cfu) per 0.2 ml. To prepare the spore inoculum, the frozen spore stock material was thawed and diluted in sterile water to achieve an inoculum shown previously to provide a specific number of spores per 0.2 ml. Mice were observed multiple times each day for 25 days after the final challenge. In preparation for passive serum transfer studies, mice were immunized with 0.1 µg of PGGA-OMPC. Serum was collected from these mice before (preimmune) and 14 days after the immunization. For the final experiment, 0.2-ml aliquots of neat sera, either preimmune or 14-day, was then mixed with 0.1 ml of saline solution aliquots containing 1.5 x 10⁴ B. anthracis Ames strain cfu for 30 min at 37 °C. Groups of 10 mice were inoculated intraperitoneally with 0.3 ml of preimmune or day-14 sera incubated with B. anthracis. A control group of 18 mice was each inoculated with 0.2 ml of saline that had been preincubated with 0.1 ml of the B. anthracis vegetative preparation. Finally, a final group of 10 mice received the same inoculum of B. anthracis incubated with 1 mg of an IgG1 anti-PGGA mAb. Mice were observed multiple times each day for a 12-day period. BALB/c mice were housed in Microisolator boxes and given water and food ad libitum. All animal studies and housing were approved and conducted following recommendations from the Institutional Animal Care and Use Committee at Merck. Within each challenge experiment, differences in survival distributions among the alternative treatment groups were assessed by the log-rank test using the LIFETEST procedure in SAS (SAS Institute Inc., Cary, NC).

	RESULTS

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Purification of PGGA—PGGA was isolated from plate-grown cultures of B. anthracis by heat extraction followed by nuclease digestion and an acid precipitation step. The yield of crude product from two independent isolations was 12 mg/plate. Trichloroacetic acid treatment proved to be a highly effective step for removing exogenous protein contaminants as the glutamic acid polypeptide remained soluble in the supernatant. The one-dimensional ¹H NMR analysis of representative crude and purified products (lot 2, see Table 1) is presented in Fig. 1. The ratio of specific PGGA concentration to gravimetric powder yielded a purity of 60–70% for crude PGGA. Unidentified contaminant bands evident in Fig. 1A were somewhat broad, suggesting that these might be due to residual protein or nucleic acid species. Ultraviolet spectroscopy of the product revealed a broad but minor absorbance centered at 263 nm along with the expected strong absorbance at 220 nm (data not shown), again suggesting the possible presence of nucleic acid. HPSEC analysis suggested that the contaminant peak was of low molecular mass and also possessed 215 nm absorbance. Hydroxyapatite chromatography was investigated as a means for removing this material. PGGA was observed to bind to the resin at high ionic strength and could be eluted with a linear gradient of increasing phosphate. The purity of fractions was assessed by analytical HPSEC, and it was found that the contaminants were quantitatively removed in the unbound flow-through fraction. Tangential flow ultrafiltration was used to remove salts and other residual low mass contaminants, and the product was stored as a freeze-dried powder. The ¹H NMR analysis of polished material (Fig. 1B) yielded powder purity of 96%. Purified PGGA was soluble in water at 3–5 mg/ml, showing a concentration-dependent viscosity that could be significantly decreased by raising the ionic strength of the medium as described previously for polyglutamate polymers (28, 29). The complete analytical profile for two representative preparations is summarized in Table 1. Most surprisingly, nucleic acid levels as measured by the Pico-Green® fluorescent dye binding were very low for both crude and purified PGGA. We showed using PAGA that the presence of high levels of polypeptide did not reduce the response of either DNA or RNA in the assay. Although Pico-Green® is typically used for DNA quantification, we found that RNA was detectable as well. The assay showed that purified PGGA preparations did not contain significant amounts of either nucleic acid. Amino acid analysis suggested that the unidentified contaminants were most likely protein or peptide fragments that co-purified through the trichloroacetic acid and dialysis steps. Although these were of low mass, it is conceivable that they were ionically bound to the polymer and removed during hydroxyapatite processing as a result of the high ionic strength of the loading buffer. The high level of purity determined by NMR was confirmed by amino acid analysis, which showed glutamic acid as the only detected residue (data not shown).

View larger version (13K): [in this window] [in a new window]   FIGURE 1. One-dimensional 1H NMR analysis of crude (A) and polished (B) PGGA. Relevant carboxylic acid proton assignments are marked. Unidentified peaks marked with asterisks are impurities removed by hydroxyapatite chromatography. Powder purity was determined with respect to internal Me2SO standard. The powder concentration was calculated according to the gravimetric weight and the solution volume in D2O. The peak integral ratio of H- over Me2SO (DMSO) was converted to a weight concentration of PGGA using the known concentration of Me2SO and the polymer repeat unit molecular weight (128 Da). The concentration ratio of PGGA from NMR to powder from gravimetric preparation yielded purity.

View larger version (17K): [in this window] [in a new window]   FIGURE 2. Structural analysis of purified PGGA. A, two-dimensional HMBC NMR spectrum of PGGA at 25 °C. The proton chemical shift is displayed horizontally, and carbon is shown vertically. The correlation between amide carbonyl carbon and H- is indicated by the shaded box. B, GC-FID analysis of PGGA. The top trace is that of purified PGGA. The middle and bottom traces are commercial enantiomerically pure L-and D-glutamic acid, respectively. C, CD analysis of purified PGGA, lots 1 and 2. Stock solutions of PGGA in water (1 mg/ml) were adjusted to 10 mM sodium phosphate, pH 7.2, and spectra were acquired at 25 °C. The polymer concentration was calculated by amino acid analysis, and the spectrum was converted to molar ellipticity as described under "Materials and Methods."

View larger version (28K): [in this window] [in a new window]   FIGURE 3. Effect of aqueous carbodiimide treatment on PGGA molecular mass and immunogenicity. Purified PGGA was treated with the indicated concentrations of EDC, pH 4.5, at room temperature as described under "Materials and Methods." A, reaction aliquots were removed at the indicated times, and the average molecular mass was determined by HPSEC/MALLS analysis. B, PGGA was treated as in A, and reaction aliquots were removed at the indicated times. Immunogenicity of the EDC-treated polymer was measured by its ability to inhibit binding of a PGGA-specific IgG1 hybridoma supernatant to ELISA plates coated with native, high molecular mass PGGA. The anti-PGGA monoclonal antibody was used at a constant dilution factor of 1:250, and inhibitor stocks were titrated from 1:2 to 1:31,250 in a 5-fold dilution series.

At a given loading ratio and reaction time, both DIPC and DMTMM afforded 3% SCL as determined by quantification of the maleimide peak, but carboxyl derivatization by isopropyl groups was measured at 5.4% for the DIPC-mediated reaction (Fig. 4A), suggesting that nonproductive adduction of reagent was occurring, possibly through formation of the stable N-acylisourea (39). In contrast, the spectrum for DMTMM-mediated coupling was much cleaner, showing only maleimide and proton resonances derived from the caproic acid spacer (Fig. 4B). HPSEC/MALLS analysis of the reaction products showed no cleavage of derivatized PAGA under these conditions (data not shown).

Next, a series of small scale PGGA activations were carried out under nonaqueous conditions using DMTMM and either EMCH or APM as the heterobifunctional activator. For this and subsequent work, the gel permeation step was eliminated, and both reagent removal and counter ion exchange were accomplished by dialysis. The results are summarized in Table 3. It was observed that the degree of activation could be adjusted by varying either the molar ratio of DMTMM relative to carboxyl or the overall reaction time. Because DMTMM was reported to be useful for amide synthesis under aqueous conditions, we also investigated this route. PGGA was solubilized in 20 mM HEPES, pH 7.3, 0.15 M NaCl buffer and reacted with either EMCH or APM in the presence of 0.5 molar eq of DMTMM. The maleimide incorporation was less than 0.5% in each case, although the amount of carboxyl derivatization was comparable with equivalent nonaqueous conditions. However, sizing analysis showed that the polymer mass had been reduced by 40-fold. Reducing the reaction time by half lowered the derivatization, but the maleimide incorporation was still low. Polymer scission also appeared to be time-dependent, because the relative mass reduction was 20–25-fold at 3 h.

View larger version (17K): [in this window] [in a new window]   FIGURE 4. PAGA activation under nonaqueous conditions. One-dimensional 1H NMR was used to assess EMCH derivatization of PAGA mediated by DIPC (A) or DMTMM (B). For reaction, PAGA was converted to the TBA+ ion form and dried. The powder was solubilized in DMF, and EMCH was added at a 1 molar eq to carboxyl groups. For the carbodiimide reaction only, 1-hydroxybenzotriazole was added at a 0.1 molar eq to carboxyl. DIPC or DMTMM was added at a 1 molar eq to carboxyl, and the reaction was allowed to proceed under N2 for 1 h on ice and then overnight at ambient temperature. Reaction products were desalted by gel permeation chromatography in DMF and dialyzed successively against 1 M NaCl and water, followed by drying. The top trace in each panel shows the regular proton spectrum, and the bottom trace is the diffusion edited spectrum. Peak assignments for PAGA protons are labeled along with the maleimide reporter peak derived from EMCH. Other nonassigned EMCH-derived shifts are labeled with asterisks. The residual isopropyl groups of the adducted DIPC are noted in A.

View larger version (21K): [in this window] [in a new window]   FIGURE 5. Dose-ranging study of response to PGGA-OMPC capsule-conjugate vaccines. Groups of 10 mice were injected intraperitoneally at 0, 14, and 28 days with PGGA-OMPC conjugate vaccines at 10 µg (•), 1.0 µg (), or 0.1 µg (), or with 10 µg of unconjugated PGGA () or 100 µg of OMPC carrier (). All formulations were adsorbed to aluminum hydroxyphosphate adjuvant. Results are expressed as the geometric mean of IgG antibody responses at the indicated times.

Mouse Immunization and Protection—A dose-ranging study (Fig. 5) was performed by immunizing groups of 10 BALB/c mice with increasing amounts of PGGA-OMPC conjugates at 2-week intervals using a 3-dose vaccination regimen. Controls received either aluminum adjuvant-adsorbed unconjugated PGGA (10 µg) or OMPC alone. The OMPC dose was matched with that in the highest dose of conjugate vaccine (50 µg). Mice responded rapidly to the first dose of PGGA-OMPC conjugate vaccines. There was little difference in response between the 10- and 1.0-µg dose levels, whereas the response to 0.1-µg dose was 10-fold lower than that elicited by the 10-µg dose level. Antibody titers rose 10-fold after the second and third doses. By contrast, mice injected with unconjugated PGGA (formulated on aluminum adjuvant) generated a modest response to the first injection that declined after subsequent doses. The results indicated that conjugation of PGGA to the potent OMPC carrier resulted in a markedly enhanced immune response.

View larger version (15K): [in this window] [in a new window]   FIGURE 6. Survival rates of immunized mice after challenge with B. anthracis vegetative bacilli. Groups of 9–10 mice from the experiment described in Fig. 5 were challenged on day 0 and on day 12 (arrows) 2 weeks after the third dose of vaccine. The number of surviving mice was assessed daily until day 25. Symbols are as in Fig. 5.

B. anthracis Challenge Studies—In a series of experiments to explore vaccine efficacy in the mouse model, mice were immunized and then challenged with either vegetative bacilli or spores. Groups of 9–10 mice from the experiment shown in Fig. 5 were challenged 2 weeks after the third dose of vaccine with 10³ cfu (day 0) and 5 x 10³ cfu (day 12) of vegetative bacilli. The number of surviving mice was assessed daily until day 25. As shown in Fig. 6, all mice immunized with 0.1–10 µg of the PGGA-OMPC conjugate vaccine survived the 10³ cfu challenge, and 100% of mice immunized with the highest dose (10 µg) of conjugate survived after the 5 x 10³ cfu challenge dose. There was no statistical evidence of a difference in survival distribution among the three PGGA-OMPC conjugate vaccine groups (p = 0.60). By contrast, 30% of mice immunized with unconjugated PGGA survived the first challenge dose, and only 20% remained alive after the second dose. Although 40% of mice immunized with OMPC survived the first challenge, none survived the second. Although the survival distributions of the unconjugated PGGA- and OMPC-only groups were not statistically different from one another (p = 0.87), they each were highly statistically different from each of the PGGA-OMPC conjugate vaccine groups (p 0.002). These results indicated that mice immunized with even the lowest dose of PGGA-OMPC conjugate vaccine were well protected from a lethal challenge with vegetative bacilli.

Additional experiments were conducted to evaluate protection against B. anthracis spore challenge. BALB/c mice were immunized on day 0 and day 14 with 0.01–1.0 µg of PGGA-OMPC conjugate vaccine, and serum was collected on days 14 and 28 for ELISA. As shown in Table 4, there was a dose-dependent antibody response to the conjugate vaccine with titers ranging from 62,602/ml to the lowest (0.01 µg) vaccine dose to 2,070,645/ml to the highest (1.0 µg) vaccine dose. Approximately 8 weeks after the second vaccine dose, all four groups of mice were challenged with a subcutaneous injection of 2 x 10⁴ B. anthracis spores. Survival was monitored for 2 weeks. Vaccination with even the lowest dose (0.01) µg of the vaccine resulted in good (80%) protection from challenge with spores from a virulent strain of B. anthracis.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
This study demonstrates for the first time that a covalently conjugated form of the native B. anthracis polypeptide capsule is able to elicit an immune response capable of protecting experimentally vaccinated animals from a lethal bacterial challenge. It has been reported that the isolated capsule is only weakly immunogenic (14, 17, 40), behaving in that respect similarly to the polysaccharide capsules of organisms such as pneumococci and meningococci (41). Recent studies employing conjugated forms of either PGGA derived from B. lichenformis (18) or synthetic poly--D-glutamic acid peptides (17) demonstrated the enhanced immunogenicity of these formulations and showed in the latter study the resultant sera capable of opsonophagocytic killing in vitro. However, neither of these demonstrated protective efficacy of the vaccine candidates.

View larger version (18K): [in this window] [in a new window]   FIGURE 7. Passive protection of mice with PGGA-specific immune serum or monoclonal antibody. Serum collected from BALB/c mice before () and 14 days post-immunization with PGGA-OMPC conjugate vaccine (•) was pooled and tested for passive protection of naive mice from B. anthracis vegetative bacilli. Neat serum was mixed at a 2:1 ratio with vegetative bacilli (104 cfu) and incubated for 30 min at 37 °C. Groups of 10 mice were then inoculated with 0.3 ml of the mixture and observed for 12 days. As described under "Materials and Methods," a separate group of 10 mice received an inoculum of 1 mg each of an IgG1 anti-PGGA mAb () that had been preincubated for 30 min with 104 vegetative bacilli. A group of 18 unimmunized control mice () were challenged with the same stock of vegetative bacilli that had been preincubated with saline for 30 min prior to challenge.

The use of HPSEC/MALLS allowed for accurate mass determinations independent of column migration behavior, and the detailed mass studies of crude and purified product were instrumental in both furthering the characterization of this antigen and in demonstrating the effect of size reduction on its immunological profile. To our knowledge, this is the first reported measurement of hydrodynamic radius for PGGA based on light scattering data. The findings are in agreement with the CD spectra and show the molecular conformation to be that of an extended random coil at neutral pH and low ionic strength, consistent with historic studies employing both CD and optical rotary dispersion analyses (36, 37, 42). Most interestingly, the hydrodynamic radius did not show any appreciable change at pH 4.5 for the control in the EDC study. Balasubramanian et al. (36) showed the ellipticity increase at 210 nm closely followed the proton titration curve of the carboxylic acid. At pH 4.5, that dissociation factor was still >0.5, indicating that, overall, the molecule was still in an ionized state. The incomplete coil-to-helix transition may account for the similar radius values. It should be noted that the optical measurement techniques yield an average conformation over the molecule and do not preclude the possible existence of shorter segments that contain a more defined structure. The study of Kimura et al. (37) demonstrated that helical transition in low mass L-polyglutamic acids (<25 kDa) followed a stepwise route, with small stretches of localized helix eventually combining to form long extended rods. It is possible that in the high mass native PGGA local environmental effects can favor and stabilize regions of secondary structure.

We have now shown definitively that aqueous carbodiimide treatment significantly reduces both molecular mass and antigenicity of the PGGA polymer. A study by Paterson and Leach (38) had demonstrated previously carbodiimide-induced cleavage of poly-L-glutamic acid at pH 5.0 in the presence or absence of ethanolamine. In discussing putative scission mechanisms, they suggested that a likely route was attack by free carboxyl groups on the O-acylisourea intermediate, basing this on the observation that cleavage was decreased in the presence of ethanolamine, which would suggest that as amide formation proceeded, fewer reactive free carboxyls were available for backbone cleavage. As their native polyglutamic acid mass was 25,000 Da, it can be reasoned that in a polymer such as PGGA, with a mass 10-fold higher, the proportion of reactive groups is significantly increased. Furthermore, at high concentrations of the carbodiimide, the potential for water-mediated conversion of the O-acylisourea back to the reactive carboxyl is greatly increased, consistent with the observed increase in scission at the higher EDC concentrations of our study. Although we do not propose a mechanism for the activation-induced cleavage of PGGA, the reaction appears to proceed readily under aqueous conditions. Carboxyl activation with EMCH in DMF did not result in chain scission using either DIPC or DMTMM, although removal of the intermediate was less efficient with the carbodiimide reagent. This lends convincing support to the postulated role for water playing a direct role in the cleavage reaction, as outlined above. Alternatively, a study of -glutamyl peptide bonds in collagen suggested side reactions leading to internal formation of glutarimide or acyl diketopiperazine ring structures in the presence of water-soluble carbodiimide. Such structures may be more susceptible to cleavage with subsequent chain scission (48). The observation that at high concentrations of EDC the mass reduction was less pronounced at pH 4.5 argues that increased protonation of the carboxyl groups has a protective effect. This would be contradictory to the putative mechanism where the protonated carboxylic acid is the reactive species. However, at the 0.5 mg/ml EDC concentration, mass reduction was more pronounced at pH 4.5. Furthermore, the degree of antibody binding was more significantly reduced for PGGA treated at pH 4.5, regardless of reagent concentration or temperature, suggesting that side chain modification may proceed without resultant polymer scission.

The observed lability of PGGA in the presence of carbodiimides has considerable implications for several recent studies in which this method of activation was employed to prepare carrier-capsule conjugates. Recently, Chabot et al. (15) reported that although native PGGA showed some efficacy against a toxin^-, capsule⁺ B. anthracis strain, the protective efficacy was completely abrogated following conjugation of the capsule to BSA using EDC at pH 4.7. This was despite the fact that the measured PGGA-specific IgG titers were increased by nearly 20-fold. This provides compelling evidence that a conformational component to the immune response may indeed be critical for protection and is consistent with our current observations that EDC-mediated reduction in capsule mass and treatment at low pH results in altered immunogenicity.

PGGA covalently coupled to OMPC by the DMTMM method described here was a highly effective immunogen, inducing polymer-specific titers in mice that were at least 3 logs higher than PGGA alone at a 100-fold lower dose. The immune response to the conjugate was observed to increase with subsequent boosting, although no increase was seen for PGGA alone post-dose 2 or 3 in the initial dose-ranging study. Most surprisingly, even a single injection of 0.01 µg of conjugated polymer provided a very robust response compared with 1.0 µg of unconjugated antigen. It is significant to note that the PGGA-OMPC conjugate raised high levels of IgG and displayed greater than 80% efficacy in both vegetative bacillus and spore challenge models, whereas unconjugated PGGA provided only 20% protection. A previous report comparing nonconjugated with BSA-conjugated PGGA found that IgM-specific titers were lower in animals vaccinated with conjugate, although the IgG titers were also significantly higher (15). However, because in this case the conjugate-immunized animals showed a complete lack of protection, the authors postulate that IgM may be the protective entity and that protective epitopes may be destroyed by the conjugation procedure. On the contrary, the positive correlation between IgG and protection demonstrated in our study clearly shows that IgG antibodies are protective. The discrepancy between the two studies can be explained by the differences in the conjugation procedures. In the previous study, the nonprotective BSA-PGGA conjugate was prepared using EDC-mediated coupling, which we have shown to significantly affect antigen mass and integrity resulting in decreased reactivity with antiserum to native PGGA. It should be recalled that in the previous study the nonconjugated PGGA was protective against a pXO1^-, pXO2⁺ strain but not wild type, whereas our conjugate vaccine showed good efficacy against the fully virulent wild type strain (pXO1⁺, pXO2⁺). Recent reports using conjugated PGGA (11, 17, 18) demonstrate a robust IgG response induced by the peptide-carrier conjugate. An antibody-mediated basis of protection is strongly supported by our demonstration of passive protection in mouse transfer experiments, as well as by previous studies that showed that conjugate-induced anti-PGGA immune sera contains opsonophagocytic activity against pXO1^-, pXO2⁺ strains of B. anthracis (17).

The ability to convert T-independent antigens such as polysaccharides and D-polypeptides to T-dependent ones capable of inducing mature IgG responses is strongly related to the choice of protein carrier (49). In this respect, OMPC offers certain advantages. First, OMPC is a component of a licensed pediatric vaccine and has a proven safety record (50). Second, in a comparison study, Donnelly et al. (51) demonstrated that in addition to the usual carrier-mediated immunoglobulin class switching, OMPC also demonstrated potent adjuvant capabilities, likely related to B-cell activation by its lipopolysaccharide component. It is postulated that this is the reason why OMPC-polysaccharide conjugates induce high serum antibody titers following one-dose immunizations, whereas other carriers typically require a three-dose regimen. The data from the present study extend that finding to D-polypeptides. Third, recent studies indicate the involvement of Toll-like receptors 2 and 4 in the immunopotentiating effects of Neisseria outer membrane protein preparations, including OMPC (52, 53). Such effects are mediated by the major membrane porins or associated lipopolysaccharide, or both, and lead to the activation of B-cells through a MyD88-dependent mechanism (52). Finally, the activation and conjugation scheme described here all