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HIV-infected individuals mount poor antibody responses to vaccines. We sought to identify the immunologic and virologic factors associated with a robust response to hepatitis Avirus (HAV) vaccine in children on highly active antiretroviral treatment.
One hundred fifty-two pediatric highly active antiretroviral treatment recipients immunized against HAV at weeks 0 and 24 had anti-HAV antibodies, CD4+, CD8+, and CD19+ cell percent assessed at weeks 0 and 32. Subgroups had HIV viremia, B- and T-cell subpopulations, and cell-mediated immunity (CMI) to HAV and other stimulants measured.
Anti-HAV antibodies after complete vaccination correlated positively with CD4+ percent and CD19+ percent and negatively with viremia and CD8+ percent at baseline, but not at 32 weeks. There were no significant correlations between anti-HAV antibodies and B- or T-cell-naïve, memory, or activated subpopulations or non-HAV CMI. Compared with children who remained HAV-CMI-negative, those who mounted HAV-CMI in response to vaccination had higher anti-HAV antibody titers and CD19+ CD21+ CD27+ memory B cell percent at 32 weeks, but no other differences.
In HIV-infected children on highly active antiretroviral treatment, control of viral replication and conserved or reconstituted CD19+ and CD4+ cell numbers and function determine a robust antibody response to anti-HAV primary immunization. Our data support a bidirectional B- and T-cell cooperation in the response to the HAV vaccine.
HIV-infected individuals have notoriously poor antibody responses to microbial antigens and vaccines, including the hepatitis A virus (HAV) vaccine.1–8 In a previous study, we showed that only a minority of children and adolescents who met AIDS immunologic criteria responded to primary anti-HAV immunization during the first year of effective highly active antiretroviral treatment (HAART).9 Others have described similarly poor responses to pneumococcal, influenza, and hepatitis B immunizations of HIV-infected individuals.1
Although HIV does not infect B cells, it decreases their function10–13 by diminishing T-cell help14 and by releasing subvirion particles that directly interfere with B-cell function. 11,12 HIV infection also has been associated with a decrease of total and memory B cells15–20 and an increase of transitional/immature B-cell subpopulations.17,21–24 In viremic patients, B cells are more susceptible to CD95L-mediated apoptosis,18 which may decrease their lifespan and account for the decrease in B cell numbers.16,25
HAART improves many aspects of the immune system of HIV-infected patients, including humoral responses to vaccines.9,25–29 However, not all HAART recipients who reconstitute or preserve their CD4+ T cells mount adequate responses to vaccines. In a previous study, we found that approximately half of the HIV-infected children and adolescents with a CD4+ T cell percentage (CD4%) ≥20% after ≥4 months of HAART had an antibody response to HAV vaccine at least one order of magnitude lower than the one reported in historical studies of healthy children and adults.30 Two years after immunization, the subjects with low antibody responses to the vaccine were more likely to lose protective anti-HAV antibody titers than those who mounted adequate antibody titers to the vaccine.
To gain more insight into the pathogenic mechanisms that govern immune responses to vaccines in HIV-infected children and to identify immunologic and virologic markers that may help optimize the timing of vaccine administration in these patients, we studied the association between humoral- and cell-mediated immune responses to the HAV vaccine and of each of these anti-HAV responses with the following variables: 1) age; 2) HIV viremia; 3) CD19+ B-cell total as well as naïve, memory, and activated subpopulations; 4) CD4+ T-cell total as well as naïve, memory, and activated subpopulations; and 5) CD8+ T-cell total as well as activated subpopulations.
HIV-infected children 2 to 21 years of age were enrolled in this Pediatric AIDS Clinical Trials Group 1008 study after obtaining local Institutional Review Board approval and informed consent. Eligibility criteria included receipt of Pneumocystis pneumoniae pneumonia prophylaxis for at least 6 months; stable antiretroviral therapy; CD4% ≥25% for children younger than 6 years and ≥20% for subjects older than 6 years; absence of previous hepatitis A vaccination or wild-type disease; and no immune globulin administration for 3 months before enrollment. Children received the first dose of HAV vaccine at enrollment and a second dose 24 weeks later. Doses were 720 and 1440 ELU of HAVRIX (Glaxo-SmithKline, Philadelphia, PA) in children younger and older than 18 years, respectively. T-cell subsets and plasma HIV RNA were measured every 8 weeks. Anti-HAV antibodies were measured at baseline and week 32, corresponding to 8 weeks after the second dose of vaccine. A subset of subjects had T-cell phenotyping and cell-mediated immunity (CMI) measured. These were randomly selected among subjects who were seronegative at baseline, received both doses of the vaccine, and had antibodies measured at week 32.
Anti-HAV antibodies were determined using a quantitative enzyme-linked immunosorbent assay. Precoated HAV microtiter plates (Viral Antigens Inc., Memphis, TN) were incubated with serial dilutions of anti-HAV World Health Organization Standards and 100 µL of serum samples diluted 1:21 in assay IgG diluent (Viral Antigens Inc.). After 1 hour of incubation, plates were washed and bound antibodies were revealed with alkaline phosphatase-conjugated goat antihuman IgG and p-NPP chromogenic substrate (Sigma, St. Louis, MO). The antibody titer of each sample was calculated by interpolating its corresponding optical density on the linear regression curve generated with the World Health Organization Standards. This assay was validated using 20 sera from HAV-naïve healthy volunteers and 10 sera with previously determined anti-HAV antibody concentrations, courtesy of Dr. Paul Willems, GlaxoSmithKline Vaccine Division. Titers ≥20 mIU/mL were considered protective against HAV infection and also defined HAV-seropositive or immune status. Responses were further classified as low if <250 mIU/mL and adequate if ≥250 mIU/mL (based on the observation that virtually all healthy individuals immunized against HAV had antibody titers >250 mIU/mL at 4 to 8 weeks after the last dose of HAV vaccine).
Peripheral blood mononuclear cells (PBMCs) were isolated from heparinized whole blood by Ficoll-Hypaque (Sigma) density gradient centrifugation. Isolated PBMCs were cryopreserved and subsequently thawed as per the International Maternal Pediatric Adolescent AIDS Clinical Trials/Pediatric AIDS Clinical Trials Group cryopreservation consensus protocol (http://impaact.s-3.com/immlab.htm).
Thawed PBMCs resusupended at a concentration of 2 × 106 cells/mL in RPMI 1640 (CellGro, Manassas, VA) with L-glutamine (Gibco, Carlsbad, CA), 1%penicillin-streptomycin, 4mM HEPES (Gibco), and 10% human AB serum (Gemini, West Sacramento, CA) were stimulated in triplicate wells with medium only, 1650 ELU/mL Hepatitis A antigen (gift of Dr. Martine Wettendorff, Glaxo SmithKline Research Laboratories, Belgium) for 6 days. PBMCs at 106 cells/mL of medium as described previously were stimulated with 2.5 µg/mL tetanus toxoid (Connaught, Swiftwater, PA), 50 µg/mL Candida albicans antigen (Greer, Lenoir, NC), 5 µg/mL phytohemagglutinin (PHA; Sigma), or no stimulant for 6 days. After stimulation, wells were pulsed with 50 µCi/well 3H-thymidine (Perkin Elmer, Waltham, MA) for 6 hours, harvested, and counted in a Packard scintillation counter. The stimulation index was calculated by dividing the median counts per minute in the stimulated wells by the median counts per minute in unstimulated wells matched by number of cells/well. A stimulation index ≥3 defined a response to HAV, candida, and tetanus and ≥10 to PHA.
For carboxyfluorosuccimide ester (CFSE) dilution assays, PBMCs were labeled with 1.5 µM CFSE (Molecular Probes, Carlsbad, CA) in 0.1% bovine serum albumin phosphate-buffered saline for 10 minutes at 37°C. Staining was quenched with cold 10% human AB serum in RPMI. An aliquot of PBMCs was removed for baseline flow analysis and the remaining cells were stimulated in tissue culture tubes with the same stimulants as described previously except for using pokeweed mitogen (Sigma) instead of PHA to elicit nonspecific proliferation. On the day of the assay, PBMC were washed in phosphate-buffered saline, stained with CD19-PE (Caltag, Carlsbad, CA) surface marker, and analyzed by flow cytometry as described subsequently.
Supernatants from HAV-stimulated PBMC 6-day cultures and unstimulated controls were used for measuring interleukin (IL)-2, IL-4, IL-5, IL-10, and interferon-γ using Luminex™ kits (Biosource, Carlsbad, CA) according to the manufacturer’s instructions. Results were expressed in pg/mL after subtracting the background cytokine concentration in unstimulated wells from that obtained in HAV-stimulated wells.
Thawed PBMCs were resuspended in phosphate-buffered saline containing 1% fetal bovine serum at a concentration of 1 × 106 cells/mL and stained with 3 or 4 combinations of the following fluorochrome-conjugated mAbs: anti-CD8-FITC, anti-CD45RA-FITC, anti-HLA-DRFITC, IgG1-FITC, anti-CD8-PE, anti-CD38-PE, anti-CD95-PE, IgG1-PE, anti-CD4-PE*Cy5, anti-CD8-PE*Cy5, anti-CD8-PE-Alexa700, or anti-CD8-APC (Pharmingen/Becton Dickinson, San Jose, CA); or with a premixed custom formulation containing anti-CD21-FITC, anti-CD27-PE, anti-CD19-PE*Alexa700, and anti-CD95-APC (Caltag Laboratories Inc./Invitrogen, Inc.). After 30 minutes of incubation at 4°C in the dark, the samples were depleted of red blood cells with FACS lysing buffer (Becton Dickinson), washed with phosphate-buffered saline, resuspended in 0.2% paraformaldehyde, and analyzed with FACS Calibur flow cytometer (Becton Dickinson) and CellQuest software (Becton Dickinson). Lymphocytes were gated based on forward versus sidescatter signals (R1) followed by either T- or B-cell selection by combining sidescatter with bright CD4, CD8, or CD19 signals (R2). T- and B-cell subsets were characterized by analyzing 5000 R1*R2 gated events and were expressed as either T- or B-cell percent positivity.
Because the immunologic outcome measures did not meet normality assumptions, nonparametric tests were used to examine the associations between antibody responses to HAV vaccine and demographic, immunologic, or virologic characteristics of the study population. The statistical significance of differences in antibody titers across different groups of subjects was assessed by Wilcoxon sum ranks (two groups) or Kruskal-Wallis tests (multiple groups). Spearman’s rank order correlation coefficients were used to examine the strength and statistical significance of associations between antibody titers and other continuous variables. When the univariate analyses identified multiple variables positively associated with the primary outcome measure, a multivariate analysis was performed using regression. Statistical significance was defined by P < 0.05.
There were 152 subjects who met inclusion criteria for this substudy by virtue of being HAV-seronegative at baseline, having received 2 doses of HAV vaccine, and being tested at 32 weeks (8 weeks after the second dose of vaccine) for antibody responses. The median age was 9 years and 13 children were between 2 and 6 years of age. Demographic characteristics such as age, sex, and race were not associated with antibody responses to the HAV vaccine (Table 1). Subjects with higher plasma HIV RNA concentrations (VL) and those with lower CD4+ T cell percentages (CD4%) at baseline had significantly lower anti-HAV antibody titers after the second dose of the HAV vaccine (P values of 0.02 and 0.04, respectively).
To identify the B-cell characteristics associated with robust antibody responses to neoantigens in HIV-infected subjects, we tested correlations between anti-HAV antibody titers at week 32, 8 weeks after the second dose of the HAV vaccine, with baseline and week 32 CD19+% (total B lymphocytes; N = 148) and subpopulations (N = 78) identified by expression of CD21, CD27, and/or CD95 (Table 2). The only parameter significantly associated with the week 32 anti-HAV antibody titers was the baseline CD19+% (N = 148, rho = 0.21, P = 0.01).
The antibody response to the HAV vaccine is T cell-dependent and, therefore, it is likely that in the context of HIV infection, anti-HAV antibody production may be limited by deficient T-cell help. To identify the T-cell characteristics associated with robust humoral immune responses to HAV in HIV-infected children on HAART, we performed correlation analyses of week 32 anti-HAV antibody titers with CD4+ and CD8+ T cell% and their subpopulations (Table 3). The anti-HAV antibody titers were weakly but statistically significantly correlated with baseline CD4+% (r = 0.16, P = 0.04) and inversely correlated with baseline CD8+% (r = 0.17; P = 0.04). Compared with CD4+ and CD8+% populations, there was a stronger inverse correlation of antibody titers with baseline CD4+ CD45RA-CD95-% central memory T cells (rho = − 0.28, P = 0.06). However, this correlation did not reach statistical significance. There were no significant correlations between antibody responses to HAV and baseline naïve or activated CD4+ or CD8+% nor with any of the T-cell phenotypic characteristics at week 32.
In HIV-infected patients, higher VLs are typically associated with lower percentages of CD4+ and B lymphocytes and higher percentages of CD8+ lymphocytes. To determine which of these variables were independently associated with the magnitude of the antibody response to HAV, we performed a multivariate regression analysis. For this analysis, the antibody titers were log transformed to meet the normal distribution criterion required for the regression analysis. Of the 152 subjects, 97 had complete sets of data, including VL, CD4+, CD8+, and CD19+%. After log transformation of the antibody titers and with the contraction of the sample size, only VL, as a discrete variable, and CD19%, as a continuous variable, remained significantly associated with the anti-HAV antibody titers (P values of 0.04 and 0.02, respectively). The backward elimination procedure applied to the regression analysis identified the CD19+% as the strongest predictor of the antibody response to HAV vaccine (P = 0.02).
To assess the relationship between antibody and CMI responses to HAV vaccine, we compared the week 32 antibody titers of HAV-CMI-positive (N = 10) with those of HAV-CMI-negative vaccinees (N = 44). Median titers were significantly higher in HAV-CMI-positive children (P = 0.02; Table 4). In contrast, there were no appreciable differences in HAV antibody titers between subjects with positive versus those with negative tetanus or Candida CMI at baseline or week 32 (Table 4), indicating that the association of HAV antibodies with HAV CMI was antigen-specific. Only 4 subjects failed to respond to PHA using a stimulation index = 10 as the threshold, which precluded an analysis of potential associations of PHA responses with anti-HAV antibody titers.
Th1 and Th2 characterization of T-cell response to HAV was attempted by measuring IL-2, IL-4, IL-5, IL-10, and interferon-g concentrations in HAV-stimulated PBMC culture supernatants. Median cytokine levels in cultures of PBMC obtained after vaccination varied from 0 pg/mL for IL-2 and IL-4 to 0.09 p/mL for IL-5, 0.47 pg/mL for IL-10, and 0.32 pg/mL for interferon-γ. These very low levels suggested that the assay, which was standardized using PBMC from healthy donors, was not sensitive enough to detect responses of HIV-infected subjects.
HAV-CMI is an important component of the protection conferred by the HAV vaccine as indicated by its correlation with antibody titers and by the fact that in the absence of seroconversion after HAV vaccination, the presence of HAVCMI is associated with protection against infection.31 To identify the factors that determine a CMI response to the HAV vaccine, we compared the demographic, virologic, and the T- and B-cell phenotypic characteristics of subjects who developed HAV-CMI at 32 weeks after vaccination versus those who did not. Of the parameters analyzed (same as for antibody responses), only the CD19+ CD21+ CD27+ memory B-cell percent at week 32 was significantly different between subjects with and without detectable HAV-CMI after vaccination: 7 HAV-CMI-positive subjects had a median CD19+ CD21+ CD27+ of 13%, whereas 49 HAV LPA-negative subjects had a median of 9% (P = 0.02, Wilcoxon signed rank test).
To examine the possibility that HAV-specific B cells proliferated in the LPA, we measured cell divisions by CFSE dye dilution coupled with CD19-surface staining in 4 HAV-vaccinated subjects with positive HAV-specific LPA. Figure 1 shows a typical representation of the results. In HAV-stimulated cultures, the CD19+ B cells represented a median (range) of 5% (0% to 17%) of the CFSElo lymphocytes, whereas corresponding percentages after pokeweed mitogen stimulation were 21% (3%to 40%). These results indicated that CD19+ B cells did not appreciably contribute to the HAV-specific LPA response.
We identified several immunologic and virologic parameters reflective of the stage of the HIV infection and/or immune reconstitution on HAART that determine the ability of HIV-infected children on HAART to mount protective antibody responses after 2 doses of HAV vaccine administered 6 months apart. In the univariate analysis, high CD19+ B lymphocyte percent, high CD4+%, undetectable VL, and low CD8+ T lymphocyte percent at the time of the administration of the first dose of HAV vaccine were associated with an antibody response similar to that of historical healthy controls,32–35 which was also necessary and sufficient to maintain protective antibody titers 2 years after vaccination in the HIV-infected children.30 In contrast, the same immunologic and virologic parameters measured concurrently with the immune response and closely after the administration of the second dose of the vaccine did not affect the magnitude of the immune response to HAV vaccine. This suggests that the primary immune response to the vaccine depended on the HAART-associated immune preservation or reconstitution, whereas the amnestic response was less affected by HIV disease-specific immunologic and virologic characteristics. Furthermore, a third dose of the HAV vaccine administered 2 years after the primary immunization generated equally high antibody titers in all vaccinees, which did not depend on CD4% or VL at any time. Others have also shown that HIV-infected adults on HAART have lower antibody responses to the first dose of HAV vaccine compared with historical controls.36 Taken together, these data support the notion that in HAART recipients, anamnestic immune responses are less likely to be affected by the stage of HIV infection than primary responses and, therefore, increasing the number of doses of vaccines may overcome a deficient primary response to the immunogens.
To establish the relative significance of the baseline immunologic and virologic characteristics with respect to antibody responses to HAV vaccine, we performed a multivariate analysis, including the parameters that were significant in the univariate analysis. The multivariate analysis included only participants with complete sets of data, which resulted in a 36% reduction of the sample size originally used to assess the relationship between lymphocyte populations and responses to HAV vaccine. After the reduction of the sample size, the association of CD4+ and CD8+ percent with antibody titers lost significance, but undetectable VL and high CD19+% at the time of the administration of the first dose of the HAV vaccine maintained their association with higher antibody responses by univariate analysis. The multivariate analysis identified the B-cell percent as the most significant factor.
We have previously documented associations between the VL and primary responses to the live attenuated varicella zoster vaccine37 or to influenza strains that were newly introduced in either the live attenuated or the inactivated seasonal influenza vaccines.38 Furthermore, this has been observed in individuals who had a diagnosis of AIDS, but also for individuals whose CD4+ cells never decreased below 20%.1,37–39 The pattern that emerges from these observations is that primary antibody responses of HIV-infected individuals on HAART are more effective when the viral replication is low or undetectable and, therefore, control of the VL should be the an important criterion for when to introduce new vaccines.
Others described associations of deficient antibody production in HIV-infected individuals with the overrepresentation of transitional/immature B-cell subpopulations expressing low CD21 and/or CD2722,23 or with the underrepresentation of memory B cells expressing adequate levels of CD21, CD27, or immunoglobulins on their surface.13,15,16,19,40 In this study, we were unable to reproduce these findings, but we identified a strong correlation between the total B-cell lymphocyte percent and the antibody responses to primary HAV immunization. Our finding suggests that for patients who start HAART after undergoing some degree of immunosuppression, it is optimal to initiate primary immunizations when their B-cell percents normalize, which generally occurs with HAART.25,26
The anti-HAV antibody production is T-cell-dependent. This may contribute to the association of CD4% at the time of the administration of the first dose of the vaccine with the antibody production. Furthermore, anti-HAV antibody titers were significantly higher in children who had a detectable HAV-specific T-cell response by LPA compared with children with negative HAV-specific LPA. This association was antigen-specific as demonstrated by the lack of association of anti-HAV antibody titers with nonspecific measures of T-cell function such as PHA, Candida, or tetanus LPA. In addition, using carboxyfluorosuccimide ester staining and flow cytometry, we found that CD4+, but not CD19+ PBMC, proliferated during the HAV-specific LPA (data not shown). Taken together, these data indicated that HAV-specific T cells boosted the antibody response to the second dose of HAV vaccine. We sought to further explore the nature of the help provided by T cells by measuring IL-2, IL-4, IL-5, IL-10, and interferon-γ secretion in LPA culture supernatants, but the cytokine levels were overall very low and there were no significant associations of Th1 or Th2 cytokines with antibody titers.
T-cell responses to HAV vaccine were found in a much lower proportion of HIV-infected children on HAART compared with historical responses in healthy HAV vaccine recipients.41 These differences are unlikely to be the result of assay conditions, because the antigens used to test both populations were obtained from the same manufacturer and other assay conditions were not appreciably different. The HAV-specific LPA differences between healthy and HIV-infected individuals are more consistent with the immune deficit of HIV-infected children that persists despite their reconstituting CD4+ cell numbers in response to HAART. We found a significant association between the CD19+ CD21+ CD27+ memory B-cell percent and positive HAV-specific LPA results. This finding underscores the importance of B- and T-cell cooperation in the response of HIV-infected children on HAART to HAV vaccine and suggests that a decrease in number or function of memory B cells may contribute to the attenuation of T-cell-mediated immune responses in HIV-infected individuals on HAART.
There was a negative association between CD8+ T-cell percent at the time of the administration of the first dose of the vaccine and antibody titers generated by the vaccine. However, there was no correlation between CD8+ T-cell subpopulations and low antibody response to the vaccine. Because CD8+ T-cell percent automatically increases with lower CD4+ and CD19+, T- and B-cell percent, respectively, it is conceivable that the association between CD8+ T cells and antibody responses to the HAV vaccine was indirect and did not reflect a pathogenic mechanism. There was also an inverse association between antibody titers and CD4+ CD45RACD95-memory T-cell percent at baseline. The significance of this correlation needs to be further explored.
In conclusion, our data indicate that antibody responses to primary immunization in HIV-infected children on HAART are primarily dependent on VL and B-cell percent and secondarily on T-cell percent. The recent World Health Organization recommendation to start HAART in all vertically infected children as soon as the diagnosis of HIV infection is established is likely to also result in improved responses to childhood vaccines, because this new recommendation should then result in administering vaccines to children with low or undetectable VL and before they experience appreciable B-cell losses.
We thank Dr. Susan Moir (National Institute of Allergy and Infectious Diseases) for scientific advice, and Dr. Jerome Bill and Rebecca Peters from the University of Colorado Denver, Eric Riedel and Nancy Raftery from the Children’s Hospital of Philadelphia, and Diane Smith from the University of California in Los Angeles for technical support. We thank the patients and their caregivers for participation. The participants were as follows: Paul Palumbo, MD, Richard Stephens, PhD, MD, Arry Dieudonne, MD, and Juliette Johnson, RN (University of Medicine and Dentistry of New Jersey/University Hospital, Newark, NJ); Abeer Khayat, MD, Saniyyah Mahmoudi, ARNP, MSN, Melissa Scites, BSN, RN, CCRC, and Sharon Cusic, RN (University of Florida Health Sciences Center, Jacksonville, FL); Vincent Bonagura, MD, Susan Schuval, MD, Constance Colter, PNP, and Lorraine Campbell, RN (Schneider Children’s Hospital, New Hyde Park, NY); Joseph A. Church, MD, Theresa M. Dunaway, RN, CCRP, MBA, and Cathy Salata, RN (Children’s Hospital Los Angeles, Los Angeles, CA); Kathleen M. Mohan, ARNP, Ann J. Melvin, MD, MPH, Lori Van Slyke, MSW, and Lisa M. Frenkel, MD (University of Washington and Children’s Hospital & Regional Medical Center of Seattle); Margaret Khoury, MD, Andrea Kovacs, MD, and James Homans, MD (University of Southern California Medical Center, Los Angeles, CA); Irma Febo, MD, Licette Lugo, MD, Ibet Heyer, RN, and Ruth Santos, RN (University of Puerto Rico, San Juan, PR); Margaret A. Keller, MD, Nasser Redjal, MD, ChrisAnna Mink, MD, and Mary Colmenar-Flores, LVN (Harbor-UCLA Medical Center, Torrance, CA); Barbara W. Stechenberg, MD, and Maripat Toye, RN, MS (Baystate Medical Center Children’s Hospital, Springfield, MA); Denise Ferraro, RN, Silvia Muniz, and Jane Perillo, MS (Stony Brook University, Stony Brook, NY); Stephen A. Spector, MD, Rolando Viani, MD, MTP, Mary Caffery, RN, and Lisa Stangl, PNP (University of California, San Diego, La Jolla, CA); Audra Deveikis, MD, and Karen Elkins, PNP (Miller Children’s Hospital, Long Beach, CA); Mary E. Paul, MD, William T. Shearer, MD, PhD, Valerie A. Nichols, BSN, RN, and Chivon D. Jackson, BSN, RN (Baylor Texas Children’s Hospital, Houston, TX); Anne Gershon, MD, Marc Foca, MD, Alice Higgins, RN, and Marie Donahue, PNP (Columbia Presbyterian Medical Center, New York, NY); Eleanor Jimenez, MD, Midnela Acevedo, MD, Milagros Gonzalez, MD, and Isos Moraima Burgos, RN (San Juan City Hospital, Rio Piedras, PR); Steven D. Douglas, MD, Richard M. Rutstein, MD, Caroll A. Vincent, CRNP, MSN, Nancy B. Tustin, MLT(ASCP)(HEW), and Donald E. Campbell, PhD (Children’s Hospital of Philadelphia, Philadelphia, PA); Elaine J. Abrams, MD, Susan Champion, MD, Delia Calo, BA, Maxine Frere, RN, and Champion, MD, MPH (Harlem Hospital, New York, NY); Hannah Gay, MD, and Sondra Sadler, RN (University of Mississippi Medical Center, Jackson, MS); Diane W. Wara, MD, Deborah Trevithick, RN, MS, Audrey Kamrin, RN, MSN, PNP, and Susan Farrales, RN, MS, FNP (University of California, San Francisco, Moffitt Hospital, San Francisco, CA); Elizabeth J. McFarland, MD, Myron J. Levin, MD, Emily Barr, PNP, and Suzanne Paul, FNP (Children’s Hospital/University of Colorado, Denver, CO); Kenneth McIntosh, MD, Nancy P. Karthas, CPNP, MS, RN, Catherine Kneut, RN, MS, CPNP, and Lynne Lewis, MS, RN, CPNP (Children’s Hospital of Boston, Boston, MA); Stephen Ira Pelton, MD, Ellen Rae Cooper, MPH, MD, and Anne Marie Regan, MEd, PNP, MSN (Boston Medical Center, Boston, MA); Ann Petru, MD, Teresa Courville, RN, MN, Karen Gold, RN, MA, and Lauren Poole, RN, FNP (Children’s Hospital, Oakland, CA); Francis Gigliotti, MD, Geoffrey Weinberg, MD, Barbra Murante, PNP, and Susan Laverty, RN (University of Rochester, Rochester, NY); Edward Handelsman, MD, Hamid Jack Moallem, MD, Denise Swindell, and JeanMarie Kaye, RN (Children’s Hospital at Downstate, Brooklyn, NY); Ellen Chadwick, MD, Debbie Cloutier, RN, Lynn Heald, RN, and Amy Talsky (Chicago Children’s Memorial Hospital, Chicago, IL); Andrew Wiznia, MD, Jacob Abadi, MD, Margaret Chin, PNP, and Karen Dorio, RN (Jacobi Medical Center, Bronx, NY); and Russell B. VanDyke, MD, Thomas Alchediak, MD, Margarita Silio, MD, and Cheryl Borned, RN (Tulane University, New Orleans, LA).
Supported by contract N01-HD-33162 (AW) and by the Colorado Center for AIDS Research Grant P30 AI054907. Overall support for the International Maternal Pediatric Adolescent AIDS Clinical Trials Group (IMPAACT) was provided by the National Institute of Allergy and Infectious Diseases (U01 AI068632) and by the Eunice Kennedy Shriver National Institute of Child Health and Human Development. This work was supported by the Statistical and Data Analysis Center at Harvard School of Public Health under the National Institute of Allergy and Infectious Diseases cooperative agreement #5 U01 AI41110 with the Pediatric AIDS Clinical Trials Group (PACTG) and #1 U01 AI068616 with the IMPAACT Group. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of Allergy and Infectious Diseases or the National Institute of Child Health and Human Development.
The authors have no conflicts of interest.