Significance Lyme disease, caused by the spirochete Borrelia burgdorferi, is the most common vector-borne disease in North America. If early infection is untreated, it can result in late-stage manifestations, including arthritis. Although antibiotics are generally effective at all stages of the disease, arthritis may persist in some patients for months to several years despite oral and intravenous antibiotic treatment. Excessive, dysregulated host immune responses are thought to play an important role in this outcome, but the underlying mechanisms are not completely understood. This study identifies the B. burgdorferi peptidoglycan, a major component of the cell wall, as an immunogen likely to contribute to inflammation during infection and in cases of postinfectious Lyme arthritis.

Abstract Lyme disease is a multisystem disorder caused by the spirochete Borrelia burgdorferi. A common late-stage complication of this disease is oligoarticular arthritis, often involving the knee. In ∼10% of cases, arthritis persists after appropriate antibiotic treatment, leading to a proliferative synovitis typical of chronic inflammatory arthritides. Here, we provide evidence that peptidoglycan (PG), a major component of the B. burgdorferi cell envelope, may contribute to the development and persistence of Lyme arthritis (LA). We show that B. burgdorferi has a chemically atypical PG (PGBb) that is not recycled during cell-wall turnover. Instead, this pathogen sheds PGBb fragments into its environment during growth. Patients with LA mount a specific immunoglobulin G response against PGBb, which is significantly higher in the synovial fluid than in the serum of the same patient. We also detect PGBb in 94% of synovial fluid samples (32 of 34) from patients with LA, many of whom had undergone oral and intravenous antibiotic treatment. These same synovial fluid samples contain proinflammatory cytokines, similar to those produced by human peripheral blood mononuclear cells stimulated with PGBb. In addition, systemic administration of PGBb in BALB/c mice elicits acute arthritis. Altogether, our study identifies PGBb as a likely contributor to inflammatory responses in LA. Persistence of this antigen in the joint may contribute to synovitis after antibiotics eradicate the pathogen. Furthermore, our finding that B. burgdorferi sheds immunogenic PGBb fragments during growth suggests a potential role for PGBb in the immunopathogenesis of other Lyme disease manifestations.

Lyme disease, caused by the spirochete Borrelia burgdorferi, is the most prevalent tick-borne human disease in temperate regions of the Northern hemisphere (1). Clinical manifestations of this disease are highly variable and can involve multiple organ systems at different times (2). Infection in humans is often heralded by a skin lesion (known as erythema migrans) at the site of the tick bite. If left untreated, the infection can disseminate to other tissues (e.g., skin, heart, central nervous system, joints) and give rise to additional skin lesions, carditis, neurological disorders, or arthritis (3⇓–5). These clinical outcomes are thought to result from host immune responses to B. burgdorferi or B. burgdorferi-derived components (6).

Arthritis is the most common late-stage clinical manifestation of Lyme disease in the United States and is often characterized by inflammation of one or more large joints (typically the knee), which are one of the sites the spirochetes frequently infiltrate (6). In ∼10% of cases, an inflammatory proliferative synovitis persists despite 2–3 mo of oral and intravenous (IV) antibiotic therapy and apparent absence of viable organisms in the synovial fluid and adjacent tissues (5, 7, 8). Development of autoimmunity is thought to contribute to the persistence of Lyme arthritis (LA), and recent studies have identified four autoantigens as targets of autoreactive T and B cell responses in patients with postinfectious LA (9⇓⇓⇓–13). It has also been proposed that B. burgdorferi-derived components may persist after initial infection and serve as immunogens, contributing to inappropriate inflammation long after the spirochetes have been killed (14). However, such persistent immunogens have yet to be identified.

B. burgdorferi does not produce lipopolysaccharides (endotoxin), and its genome does not appear to encode effectors that might act as toxins (15, 16). Therefore, most studies to date have focused on surface-exposed lipoproteins anchored in the outer membrane of B. burgdorferi. These lipoproteins play important roles in various aspects of tick colonization, mammalian infection, and host immune evasion and response (17⇓–19). Comparatively, the peptidoglycan (PG), an essential component of bacterial cell envelopes, has received very little attention. The PG, which is made of glycan strands cross-linked by short peptides, forms a polymeric meshwork around the cytoplasmic membrane and provides resistance against intracellular osmotic pressure (20, 21). PG is also a microbe-associated molecular pattern that can stimulate innate immune pathways in animals, resulting in inflammation (22). PG from Gram-positive bacteria administered intraarticularly or systemically can induce acute arthritis in mice and rats (23⇓⇓⇓⇓⇓–29). NOD2, an innate immunity protein recognizing a PG moiety, has been implicated in proinflammatory cytokine production and immune tolerance during B. burgdorferi infection in mice (30, 31). Furthermore, a 1990 report has shown that B. burgdorferi PG (PGBb) stimulates interleukin 1 (IL-1) production in macrophages in vitro and that intradermal injection of PGBb in human volunteers results in skin reactions characteristic of inflammation (32). Despite these observations, a potential role for PGBb in B. burgdorferi pathogenesis has not been directly examined.

In diderm bacteria, including B. burgdorferi, the outer membrane shields the PG meshwork from the external environment. Exposure of PGBb to the host immune system may, however, still be significant for two reasons. First, spirochete death, which occurs during early stages of transmission and dissemination (33), may result in PGBb exposure to host immune cells. Second, sequence homology analyses predict that B. burgdorferi lacks a PG recycling pathway (34). Absence of PG recycling suggests that large amounts of PG fragments (known as muropeptides) may be released into the host environment during spirochetal growth. Bacteria degrade ∼40–50% of their PG per generation, as part of the normal PG remodeling process required for cell wall expansion (34⇓–36). In Gram-negative/diderm bacteria, the vast majority of muropeptides produced during normal PG turnover are typically recycled. During this process, muropeptides are transported into the cytoplasm by an inner membrane permease (AmpG), processed by PG recycling proteins (e.g., AmpD and LdcA), and reincorporated into the PG biosynthetic pathway for reuse (SI Appendix, Fig. S1A) (34). Bacterial mutants that lack AmpG shed a large amount of muropeptides into their environment during growth (SI Appendix, Fig. S1B) (36⇓⇓–39). The apparent absence of a canonical muropeptide recycling pathway in B. burgdorferi suggests the possibility that muropeptides produced during normal PG turnover may be released into the extracellular milieu where the host immune system would be able to detect them. These considerations motivated us to test the hypothesis that PGBb is an antigen contributing to proinflammatory responses during the infectious and postinfectious phases of LA.

Methods Bacterial Strains, Cell Lines, and Growth Conditions. A clone of the B. burgdorferi type strain B31 (MI) (16) was used in all experiments involving this bacterium. Other bacteria used in this study include S. aureus SA113, B. subtilis 168, and E. coli K-12 MG1655. Unless otherwise noted, B. burgdorferi was cultured at 34 °C in complete BSK II medium containing 6% rabbit serum (78). All other bacteria were grown at 37 °C in LB medium. HEK 293-derived human NOD1 and NOD2 reporter cell lines (InvivoGen) were cultured at 37 °C under 5% CO 2 in RPMI medium containing 10% (vol/vol) FBS and blasticidin S (30 µg/mL), Zeocin (100 µg/mL), and Normocin (100 µg/mL). Fresh PBMCs from healthy human subjects were obtained from mixed donor samples (Zen-Bio) and used in assays in the recommended PBMC culture medium (Zen-Bio). PG Purification. PGBb was purified as described previously (79), which is an adaptation of the Glauner protocol (80). For immunological and mouse studies, a few modifications were made to increase yield and ensure purity. PGBb was purified from 2–3 L of B. burgdorferi culture. Before protease treatment with 300 µg/mL α-chymotrypsin (Sigma-Aldrich), insoluble PGBb was treated with 50 U of DNase (Zymogen) and 10 U of RNase A (Promega) for 2 h, followed by a 2-h treatment with 10 µg/mL amylase (Sigma-Aldrich). After protease digestion, PGBb sacculi were harvested and washed three times with 10 mL endotoxin-free water, once with 10 mL 0.5 M EDTA, and three more times with water. A similar procedure was performed to purify PG from E. coli. For PG preparations from Gram-positive bacteria, the cell walls were broken using a kit (Precellys Microorganism Lysing Kit) that includes 7-mL tubes containing glass beads before sodium dodecyl sulfate (SDS) solubilization and enzymatic treatment. The Precellys Evolution homogenizer was set to 10 cycles of 30 s at 8,500 rpm with a 60-s rest period between each cycle. Afterward, samples were treated with 48% hydrofluoric acid for 48 h at 4 °C to hydrolyze PG-bound teichoic acids as previously described (81). Post hydrolysis, PG sacculi were harvested and washed as described here earlier. The concentration of all purified PG preparations was determined by dry weight and confirmed by SLP assay as previously described (82). PG Structural and Chemical Analysis. Purified PGBb (∼100 µg) was digested with cellosyl (25 µg/mL) for 14–16 h at 37 °C, and the resulting muropeptides were analyzed by LC-MS as reported previously (83). For the chemical analysis, purified PGBb (0.7 mg) was hydrolyzed (200 µL 4 N HCl, 100 °C, 16 h) in a sealed ampoule. The hydrolysate was evaporated to dryness in a gentle stream of air at 60 °C. The residue was dissolved in 200 µL water and dried down again to remove residual HCl. The amino acids of the hydrolysate were transformed into N-pentafluoropropionyl amino acid isopropylesters according to protocol 11 described in a previous review (84). These amino acid derivatives were analyzed by GC (GC-14A; Shimadzu) with a CP-ChiraSil-l-Val column (Agilent Technologies, CP495) following protocol 11 and by GC-MS using a 320 Single Quad instrument (Varian) equipped with a VF-5ms column (CP8944; Agilent Technologies) using protocol 10 (84). To verify the incorporation of l-Orn into the PGBb (SI Appendix, Fig. S2B), B. burgdorferi was cultured in 500 mL complete BSK II medium to a density of 106 cells per milliliter. Cells were harvested by centrifugation (3,500 × g for 20 min) and resuspended in 50 mL of prewarmed, modified medium (25% BSK II in PBS plus 1.2% rabbit serum) (85) containing 7.5 µCi/mL of 3H l-Orn (Perkin-Elmer). After 48 h of incubation, unincorporated radiolabeled l-Orn was removed by centrifugation (3,500 × g for 20 min) and three washes with 40 mL of PBS. After each wash, cells were harvested by centrifugation at 3,500 × g for 10 min. After the washes, the cells were gently resuspended in 5 mL of PBS and PG was purified as described earlier. PG Turnover Studies. To track the turnover of PGBb over time (Fig. 1B), we used two different protocols. In the first one, 500 mL culture of B. burgdorferi at a cell density of 106 cells per milliliter was pulse-labeled with 7.5 µCi/mL of 14C l-Orn as described earlier. After three washes and centrifugation, cells were gently resuspended to a final concentration of 5 × 104 cells per milliliter in 250 mL prewarmed BSK II complete medium [which includes rabbit serum that contains Orn (86)]. Retention of radiolabel into the PGBb was tracked by removing a 25-mL culture volume at various time points, harvesting the cells by centrifugation (3,500 × g for 20 min), washing cells once with 25 mL of PBS, and harvesting cells at 3,500 × g for 10 min. Pelleted cells were resuspended, and cells were solubilized in a boiling solution of 4% SDS for 30 min. SDS-insoluble PGBb was pelleted at 145,000 × g for ∼30 min and analyzed by liquid scintillation. In the second protocol, a 250-mL culture of B. burgdorferi at 106 cells per milliliter was centrifuged (4,000 × g for 20 min) and cells were resuspended in 50 mL of prewarmed, modified medium (as described earlier) containing 7.5 µCi/mL of 3H l-Orn. After 48 h of incubation, unincorporated radiolabeled l-Orn was removed by centrifugation (4,000 × g for 20 min) and three washes with 40 mL of PBS. After each wash, cells were harvested by centrifugation at 3,000 × g for 10 min. After washes, cells were resuspended in 125 mL of BSK II complete medium at a density of 5 × 104 cells per milliliter. The remaining steps were the same as the first protocol except that 10 mL of culture was removed at each time point and cells were harvested at 4,000 × g for 20 min. Both protocols gave highly similar results (Fig. 1B). NOD Activation Assay. Time-course experiments to monitor the release of muropeptides were performed to ensure that potential stress during the radiolabeling procedure (as detailed earlier) or washes did not significantly alter our findings. In these experiments, 10 mL of culture was removed from a 250-mL batch culture, cells were enumerated, and 8 mL of culture was filtered by using a 0.1-µm filter under a vacuum. From the filtered flow-through, 5 mL was processed through a YM-3 Amicon filter to selectively exclude biomolecules greater than 3,000 Da. Column flow-through (4 mL) was lyophilized and resuspended in 1 mL of endotoxin-free Dulbecco’s PBS (DPBS), resulting in a 4× solution of the culture supernatant. Sterile BSK II complete medium (without phenol red) was processed similarly to serve as control medium to which each signal was background-subtracted. HEK-Blue hNOD1 and hNOD2 cells were cultured to 60–70% confluence, washed with PBS, enumerated, and resuspended in QUANTI-Blue detection medium (InvivoGen) at a final concentration of 2.5 × 105 cells per milliliter. HEK-Blue hNOD1 or hNOD2 cells (180 µL per well) were incubated in 96-well plate in triplicate with 20 µL of a three-time dilution (in DPBS) of the 4× culture-supernatant solution. Cells were incubated at 37 °C in 5% CO 2 for 18 h. Colorimetric quantification of NF-κB activity through NOD1 or NOD2 activation was measured at 650 nm. Gefitinib (Sigma), an inhibitor that interferes with adaptor protein RIP2 signaling (43), was used at a final concentration of 20 µM. Human Subject Samples. All work with human samples was approved by the human investigations committee at Massachusetts General Hospital granted to A. Steere. Patients with Lyme disease satisfied the criteria put forth by the Centers for Disease Control and Prevention (87). Patients with LA were treated with 1–2 mo of oral antibiotic therapy (usually doxycycline), followed by an additional 1 mo of IV antibiotic therapy (ceftriaxone) if needed, as described by the Infectious Diseases Society of America (88). Control synovial fluid samples were acquired from patients with rheumatoid arthritis, psoriatic arthritis, and osteoarthritis who met the criteria associated with each disease (89⇓–91). Serum and synovial fluid samples were collected and then centrifuged at 300 × g for 10 min, followed by another centrifugation at 3,000 × g for another 10 min to remove cells and cell debris as previously described (92). All samples were stored at −80 °C and did not undergo more than two freeze–thaw cycles. PCR Analysis. Serum and synovial fluid samples were screened by PCR for amplification of the B. burgdorferi flaB gene by using the fla-3 (5′-GGGTCTCAAGCGTCTTGG-3′) and fla-4 (5′-GAACCGGTGCAGCCTGAG-3′) oligonucleotides and Phusion Polymerase (New England Biolabs). The cycling conditions were as follows: 1 cycle at 98 °C for 30 s and 45 cycles of 98 °C for 12 s, 58 °C for 20 s, and 70 °C for 15 s, followed by a final extension at 70 °C for 5 min. All reactions were subjected to DNA agarose electrophoresis and visualized by ethidium bromide staining. Visible products were apparent for serum samples 9, 13, 17, 20, and 33 and for synovial fluid samples 9, 13, 17, and 20 (SI Appendix, Fig. S4C). ELISA. To quantify the level of anti-PG IgG in patient samples, purified PG sacculi (100 µg/mL) in PBS with 0.01% SDS were immobilized on poly-lysine–coated microtiter plates overnight at 4 °C. Unbound material was removed through three washes with PBS-T (PBS plus 0.05% Tween 20). The wells were then “blocked” for 2 h at 37 °C using SEA-BLOCK (Thermo Fisher Scientific). Serum and synovial fluid samples were diluted 1:25 in PBS and incubated with substrates (or diluent control) for 2 h at room temperature with gentle rocking. Unbound material was washed with PBS-T. After washing, plates were incubated with anti-human IgG-HRP (1:25,000; Sigma-Aldrich), and bound IgG was detected by using 1-step Turbo TMB substrate (Thermo Fisher Scientific). Reported IgG response was determined by measuring absorbance at 450 nm, following background subtraction for each patient sample signal attained in the absence of PG ligand. Detection of PGBb in patient samples was performed by using a competitive ELISA and rabbit anti-PGBb polyclonal antibodies produced as a fee-for-service by Cocalico Biologicals (Thermo Fisher Scientific). Briefly, purified PGBb (0.5 mg/mL) was used to immunize two New Zealand White rabbits according to protocols approved by the animal care and use committee of Cocalico Biologicals. After one dose, two boosters of 0.5 mg/mL were administered 1 wk apart. Serum from blood samples collected on days 53 and 54 was assayed for PGBb specificity by competitive ELISA. Competitive ELISA involved coating plates with 100 µg/mL of PGBb as described earlier. Rabbit serum containing anti-PGBb IgG was diluted 1:350 in PBS and preincubated for 2 h with titrating amounts (10 ng/mL to 10 pg/mL) of different bacterial PG preparations with gentle mixing at room temperature before 1 h incubation with PGBb-coated plates. All patient samples were diluted 1:5 in PBS and otherwise treated exactly as the PGBb standards of known concentration. Rabbit anti-IgG-HRP (Bio-Rad) diluted 1:3,000 was used to detect anti-PGBb. Standard curves were created by using 1/absorbance values (at 450 nm) produced with known concentrations of each PG preparation. Data were fitted by a third-order polynomial equation. Standard curve experiments were performed on the same day as the serum and synovial fluid analyses and used to back-calculate the amount of PGBb in each patient sample. PBMC Stimulation and Cytokine Analysis. Muropeptides were generated by digesting 1 mL of purified PG (120 µg/mL) with Streptomyces globisporus mutanolysin (1,000 U/mL; Sigma-Aldrich) for 4 h at 37 °C in buffer (50 mM MES, 1 mM MgCl 2 , pH 6), followed by another incubation of mutanolysin (∼500 U) overnight at 37 °C. Undigested material was harvested by centrifugation at 150,000 × g for 30 min at 12 °C. The soluble muropeptides were lyophilized, and their amount was determined by weight. Upon arrival, fresh PBMCs (Zen-Bio) were seeded in 12-well plates at 106 cells per milliliter and allowed to rest at 37 °C under 5% CO 2 atmosphere for 24 h before further manipulation. After stimulation with 100 µg/mL of digested or polymeric PG, cells were harvested by centrifugation at 600 × g for 8 min and supernatants were collected and stored at −80 °C for further analysis. All cytokines were assayed using Luminex bead arrays (Agilent) following the manufacturer’s recommendations. All supernatants were diluted 1:5 in PBS and analyzed in duplicate. Serum and synovial fluid samples from patients with LA, diluted 1:3 in PBS, were similarly analyzed in duplicate and run on the same day as the PBMC supernatants. The concentration of cytokines (in picograms per milliliter) from patient samples were log 2 -transformed to create the heat map (SI Appendix, Fig. S6). PG Injection in Mice and Histopathology. Purified PGBb was lyophilized, weighed, and resuspended to a final concentration of 2 μg/μL in DPBS. To achieve even dispersal PGBb in DPBS, the suspension was subjected to four rounds of sonication (15 s each) on ice using a Branson Digital Sonifier set to 45% amplitude. Fragmented PGBb (100 µL, i.e., 200 μg PGBb) was administered IV to each of 12 female BALB/c mice (5–6 wk old) by tail vein injection. In parallel, 12 BALB/c mice (age- and sex-matched) were injected IV with 100 μL DPBS. All mice were then examined daily for foot and ankle swelling and assigned a clinical arthritis score as previously described (29). Briefly, arthritis scores were computed by summing the individual scores for both hind paws, each graded as follows: 0, normal paw, no redness or swelling; 1, some swelling of ankle; 2, moderate swelling and redness of ankle; 3, moderate swelling and redness of ankle and some swelling of foot pad and/or digits; and 4, pronounced swelling and redness of the whole paw. Each group of mice was also evaluated for the prevalence of arthritis (defined as the percentage of mice with an arthritis score of at least 1). Half of the mice in each group (n = 6) were euthanized by CO 2 asphyxiation on days 2 and 4 postinjection, and both hind limbs from each animal were immediately fixed in 10% formalin and subsequently decalcified, embedded in paraffin, sectioned, and stained with hematoxylin-eosin by routine methods. For each mouse, one stained section per hind limb midlevel (to include the stifle and tibiotarsal joints) was analyzed. Sections were analyzed, and stifle and tibiotarsal inflammation was scored blindly by a veterinarian (C.J.B.) formally trained in pathology with years of experience in scoring mice for inflammation using previously published criteria (93). All procedures involving mice were approved by the Yale University Institutional Animal Care and Use Committee.

Acknowledgments We thank Alexia Belperron, Jialing Mao, and Nicole D’Angelo for assistance with the mouse experiments; Roman Dziarski for advice in planning the PG injection studies; and Dr. Patricia Rosa and the laboratories of C.J.‐W. and B.L.J. for valuable discussions and critical reading of the manuscript. This study was supported in part by Wellcome Trust grant 101824/Z/13/Z (to W.V.) and National Institutes of Health grants AI101175 and AI144365. C.J.-W. is an investigator of the Howard Hughes Medical Institute.

Footnotes Author contributions: B.L.J. and C.J.-W. designed research; B.L.J., R.B.L., Z.A.K., J.B., K.S., C.J.B., J.G., P.S., W.V., L.K.B., and A.C.S. performed research; B.L.J., R.B.L., Z.A.K., S.K.G., and W.V. analyzed data; and B.L.J., Z.A.K., and C.J.-W. wrote the paper with assistance from all authors.

Reviewers: T.G.B., Harvard Medical School; and J.D.R., University of Connecticut Health Sciences Center.

The authors declare no conflict of interest.

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