Approval for using residual material from routinely collected peripheral blood specimens from healthy normal blood donors was obtained from the Institutional Review Board at Baystate Medical Center. De-identified NBD residuals collected in ethylenediaminetetraacetic acid (EDTA) tubes were obtained from the Blood Bank at Baystate Medical Center in Springfield, MA from November 2001 to October 2002. Samples were collected monthly and stored at 4°C until tested. For other assessments, de-identified fresh blood sample residuals (FB), collected in EDTA tubes, were obtained on a weekly basis from patients seen at the University Health Services (UHS) and used immediately. Unless otherwise specified, conjugated secondary antibody reagents were obtained from Jackson ImmunoResearch Laboratories, West Grove PA; the standard incubation was 1 h at room temperature (RT) unless otherwise specified.

A 14 amino acid C. pneumoniae specific peptide sequence deriving from the gene encoding the major outer membrane porin protein of C. pneumoniae AR 39 30 was synthesized (Sigma Genosys, The Woodlands, TX). It served as the antigen in ELISA tests for C. pneumoniae specific antibody in NBD plasma samples. ELISA wells (NUNC™, Nunc A/S, Roskilde, Denmark) were coated for 1 h with 100 ul containing 1 μg/ml peptide in phosphate buffered saline pH 7.2 (PBS). Plasma samples were serially diluted in ELISA wells using 0.1% BSA in PBS and standard protocols for blocking (3% bovine serum albumin: BSA, Sigma, St. Louis, MO), incubations, and washes (X3 with 0.1%BSA/PBS). Detection of bound human antibody used alkaline phosphatase (AP)-conjugated goat anti-human IgG and after 45 min incubation and multiple rinses, 100 μl/well of P-Nitrophenyl Phosphate substrate (pNPP, Sigma, St. Louis, MO) was added. Absorbances at 405 nm were measured with a VERSAmax Microplate reader (Molecular Devices Sunny Vale, CA). Titers were determined by comparison of absorbance readings of known positive and negative controls with those of the test samples.

Smears prepared with ~20 μl/ slide of whole blood collected in EDTA were air dried then fixed 10 min in 70% cold methanol (MEOH). Following PBS rinses, slides were incubated in a moist chamber with a rabbit or guinea pig anti-Chlamydia EB serum (Biomeda Corp. Foster City, CA) then multiply rinsed in PBS. Bound anti-Chlamydia antibody was detected with fluoresein isothiocynate (FITC)-conjugated goat anti-rabbit or donkey anti-guinea pig IgG. Primary and secondary antibody incubations used the standard protocol noted above. After PBS rinsing X3, coverslips were mounted using Fluoromount G (BioWhittaker, Walkersville, MD), sealed, then examined and photographed using a Nikon Eclipse E600 epifluorescence microscope and a SPOT digital camera (Digital Instruments Inc.) or a Zeiss LSM 510 Meta Confocal System

The buffy coat (BC) containing WBC, was removed from each 5 ml NBD sample, rinsed X4 with sterile PBS, treated to release EBs and cultured on J774A.1 monolayers as described previously 30. At 96 h post infection (pi) monolayers on coverslips were rinsed with PBS, fixed with 70% MEOH, rinsed again and immunostained with a rabbit anti-Chlamydia EB, and then mounted as noted above.

Smears of BC cell pellets from each 5 ml fresh blood samples (FB), average donor age 28 years, were fixed with 70% MEOH. Except as specified, immunostaining used rabbit polyclonal anti-Chlamydia EB antibody followed by a TRITC conjugated goat anti-rabbit antibody. Specific cell type detection used mouse monoclonals: anti-Human CD14 (0.2 μg/10⁶cells) (Sigma, Saint Louis, MO), or anti-Human CD16b (1.0 μg/10⁶cells) (BD Biosciences Pharmigen, San Diego, CA.). Binding was detected with FITC-conjugated goat anti-mouse IgG. For samples incubated with PE-conjugated anti-Human CDw125 (1.0 μg/10⁶cells), bound anti-Chlamydia antibody detection used FITC conjugated anti-rabbit antibody. Slides were incubated, rinsed, mounted and sealed as described above and digital images of optical sections through these samples were acquired with a Zeiss LSM 510 Meta Confocal System.

Flow cytometry (FC) quantification of Chlamydia-infected peripheral blood WBC used PBS rinsed BC cells from smear positive NBD samples. The WBC pool was fixed and permeabilized (1% paraformaldehyde and 1% Triton X-100, 10 min. at RT; Aldrich Chemical Company, Inc., Milwaukee, WI). PBS rinsed RBCs from the same samples were also pooled. BC and RBC were separately incubated with pre-diluted guinea pig anti-Chlamydia primary antibody followed by incubation with PE-conjugated F(ab')₂ donkey anti-guinea pig IgG, PBS rinsed X3, and mono-dispersed using a nylon mesh filter (Lab-Line Instruments Inc, Melrose Park, IL). A WBC dilution series was prepared in separate tubes as follows: 100%, 90%, 80%, 70%, 60%, 50%, 40%, 30%, 20%, 10%, 5%, 2%, 1%, 0.5%, and 0% WBC/tube. Sufficient RBCs were added to obtain a total of 50,000 cells/tube and the series analyzed by FACScan (Becton, Dickinson and Company, Franklin Lakes, NJ).

BC isolated from FB samples was pre-screened for Chlamydia infected cells by anti-Chlamydia immunostaining as described above. BC from four smear positive samples and separately, BC from four smear negative samples were pooled. Each pool was washed, fixed and permeabilized as described above. After counting, aliquots of the cell pools were incubated with the polyclonal guinea pig anti-Chlamydia primary antibody and one of the three monoclonal antibodies, at the concentrations noted above. To detect Chlamydia and CD14 or CD16b, BC were incubated with PE-conjugated F(ab')₂ donkey anti-guinea pig IgG and FITC-conjugated rabbit anti-mouse IgG. Chlamydia in BC immunostained with PE-conjugated anti-CDw125, were detected with FITC conjugated F(ab')₂ donkey anti-guinea pig IgG. After standard incubations and rinsing, samples were mono-dispersed, counted, diluted to obtain 1 × 10⁶ cells/ml of sample and analyzed using the FACScan instrument.

NBD sample data were analyzed using the SPSS^® 11.5 graduate pack statistics program [SPSS, Inc., Chicago, IL]. Comparisons of culture and smear positivity used Cross-tabs with the Fisher exact test to uncover nonrandom associations between these two categorical variables and Chi-Squared test and Kappa measure of agreement to determine significance. To verify the reproducibility of culture and smear assessments all samples were initially tested in duplicate. Subsequently, approximately 20% of the samples were re-tested by each method. The SD and CV were calculated based on the differences in the positive and negative percentages determined by the original vs. the repeat tests. All tests were two sided, and the level of significance was p ≤ 0.05.

A random cohort of blood donor sample residuals was obtained from the Blood Bank unit at Baystate Medical Center over a 12 month period. Donor demographics are presented in Table 1. As shown, the average age of this cohort was 44 years and consisted of 55% male and 45% female donors. Plasma was removed from the samples, and an aliquot assayed by ELISA for anti-C. pneumoniae antibody. As Table 2 shows, 219 (47.7%) of 459 samples tested exhibited a positive titer against the 14 amino acid C. pneumoniae-specific peptide. This level of sero-positive status is fully consistent with published reports for this age group and indicates this cohort is not anomalous in this respect.

Samples, including blood, are routinely assessed for Chlamydia by PCR and RT-PCR 5131415 but inherently PCR cannot demonstrate whether this DNA represents fragments or viable, infectious organisms 13141516 and immunostaining to detect infected PBMCs is not routinely used. This study therefore included immunofluorescent staining as described above to identify Chlamydia-infected NBD cells. As Figure 1A-C shows, Chlamydia positive material can be readily detected within lymphocyte-like and monocyte-like cells and examination of all 459 samples demonstrated 113 (24.6%) were positive for Chlamydia within peripheral WBCs (Table 2). The blood residuals, normally ≥30/month, were obtained over a 12 month period (Table 3) and comparison by month indicated that samples collected in December exhibited distinctly higher carriage of intracellular Chlamydia: 42% smear positive and 33% culture positive. In contrast, for August and November carriage was notably lower. Culture and smear positive results for these two months were 15%, 15% and 17%, 12% respectively. These divergent percentages may reflect a variable carriage rate in this cohort during the year since the sample numbers tested for each of these months were similar: 34, 35 and 39, respectively.

Immunostain for Chlamydia antigens demonstrated their presence within PBMC. Therefore we next assessed whether infectious organisms per se are present. Chlamydiae are known to infect macrophages in vivo 171819 and in vitro the J774A.1 macrophage monolayers are readily infected by Chlamydia and support growth for ~120 h pi. Using an effective culture method we developed previously and have described 30313233, WBC lysates were prepared and tested in duplicate on J774A.1 monolayers. At 96 h pi the monolayers were rinsed, fixed, and immunostained to detect Chlamydia. The results demonstrated that of the 459 samples, 114 (24.8%) were positive for growth of Chlamydia (Table 2). Figure 2 shows representative immunostained monolayers incubated with WBC derived inoculum from Chlamydia smear positive (Figure 2A) or negative samples (Figure 2B). At 96 h pi the cultures of smear positive samples contain large, late-stage inclusions as well as small, newly forming inclusions and released extracellular EBs. Table 2 shows that of the 459 samples tested 76 were positive by both smear and culture tests. Of the total positive samples, 37 were positive by smear but negative for cultivable organisms, potentially an indicator of persistence. Conversely 38 were negative by smear and positive by culture; this last finding may reflect the small sample size (20 μl) used in smear preparations. Together however the smear and in vitro culture findings demonstrated that not only do NBD samples carry PBMC infected by Chlamydia and visually identifiable by fluorescent microscopy, but frequently these WBC borne chlamydiae are infectious.

Despite the demonstration of Chlamydia DNA in PBMC samples, and demonstrations that in vitro numerous cell lines can be infected by different Chlamydia species and serovars, the specific types of WBC infected in vivo have not been fully examined. We selected three specific CD markers to identify WBC types and used them in conjunction with anti-Chlamydia antibody to immunostain WBCs in peripheral blood sample residuals. Since WBC inherently exhibit different life spans and can exhibit differential fragility, these assessments utilized fresh blood sample residuals. Smears of these FB samples were prepared, fixed and dual immunostained to detect Chlamydia and either CD14 (monocytes), CDw125 (eosinophils/basophils) or CD16b (neutrophil). Using Meta Confocal microscopy, optical sections through cells were examined and digital images acquired. Figure 3 shows representative examples of Chlamydia-infected monocyte (CD14), eosinophil/basophil (CDw125) and neutrophil (CD16b) cells. These can exhibit punctate fluorescent staining (row CD14 and CD125w) that reflects the expected uneven distribution of antigens. This is especially evident in non adherent cells and readily detectible in an optical section through a chlamydial inclusion. The anti-Chlamydia column shows the presence and distribution of Chlamydia positive material within the individual optical section, while the CD marker column identifies the cell type specific marker. The yellow or orange/yellow merge demonstrates the co-localization of the immunoprobes in that same optical section. The final column displays the DIC images and shows the position and general morphology of the individual cells. A comparison of the DIC and Merge images of the same row shows additional cells can be present that are not immunostained by either antibody (Figure 3-CD14 and CDw125). This observation serves to indicate the specificity of the immunoprobes. Figure 4 presents merged images obtained at three different levels from the same Z-axis optical sections through the cells shown in Figure 3. As the optical sections in the CD16b series show, the CD16b protein co-associates with the chlamydial antigens (6 μ) but also is present at the cell surface (10 μ).

Immunofluorescence allows detection of chlamydial antigens by microscopy but smears are inherently susceptible to significant sampling error. They also can not support quantification of total Chlamydia infected cell numbers in peripheral blood samples. We therefore utilized FC to quantify potentially 'sticky' Chlamydia infected BC samples. After pooling WBC pre-screened by immunostain for intracellular Chlamydia, a dilution series was prepared and FACScan assessments for each dilution analyzed (Figure 5). The data shows the series is linear over a broad range of infected cell numbers and that numeric differences between dilutions in the series are readily evident. As Figure 5 also shows, even in the most dilute sample that contains only 0.5% WBC, 3 ± 1.5 Chlamydia positive cells above background were detectible in the 10,000 cell sample. In addition to indicating the utility of FC for detecting Chlamydia in peripheral WBC samples, the series also serves as a control for subsequent FC directed towards quantifying specific WBC types infected by Chlamydia in vivo.

Given the demonstrated linearity and sensitivity, we utilized dual channel FC as a rapid, sensitive tool to quantify both the infected cell load (ICL) and the specific WBC types infected in vivo. Using fresh blood samples, two WBC pools were prepared and immunostained. One pool contained WBC from samples positive by blood smear anti-Chlamydia immunostain examination. The second pool, termed weakly positive, was prepared from samples Chlamydia negative by immunostain but positive by initial FC assessment. Aliquots of BC also were immunostained with a single antibody and used to establish the appropriate compensation for signal overlap between the two fluorescence channels and to establish appropriate, conservative demarcations of fluorescence positive.

Peripheral WBC populations contain neutrophils (CD16b), monocytes (CD14) and eosinophil/basophils (CDw125) typically present at levels of 60–70%, 3–8%, and 1–4% respectively. Aliquots of each WBC pool were dual immunostained with anti-Chlamydia antibody and monoclonal antibody specific for CD16b, CD14, or CDw125 as described above. The dual channel FC data (Figure 6) indicate that in the population of 10,000 cells counted for the weakly positive pool, 10.4% were positive for both Chlamydia and the neutrophil marker, CD16b, while in the positive pool 7.4% exhibited dual positive fluorescence. Notably, the dot plot for the positive pool also contains a distinct distribution ('cloud') of cells lying somewhat above the CD16b line of demarcation that clearly is absent from the dot plot of the weakly positive sample. In this 'cloud' region, numerous cells in the positive sample exhibit a lower level of CD16b expression, as indicated by their location relative to the CD16b positive demarcation line. This low CD16b expression level would be consistent with the cells being immature neutrophils. Dual immunostaining of Chlamydia-positive monocytes (CD14) demonstrated that in the weakly positive pool, 2.3% of the 10,000 cells was dual positive, while in the positive pool, 4.1% of cells, nearly double, was dual positive. Finally, in aliquots immunostained to detect Chlamydia infected eosinophil/basophils (CDw125), dual positives were identified in 1.9% of cells (weakly positive sample) and 5.4% (positive sample), more than double the number of cells.

This quantitative dual channel FC data indicates that in vivo all three of these cell types can become infected, and is consistent with our identification in dual immunostained smears. The data also shows FC can quantify each cell type and that both the dot plot patterns and cell type specific numbers clearly differ for the weakly positive vs. positive samples. Therefore even at the minimal level of Chlamydia-infected cell carriage in these samples, cells of each type become infected in vivo and both the total infected cell load (ICL) and the infected cell type specific load (ICSL) can be quantified.