Breast Milk-Derived Antigen-Specific CD8+ T Cells: An Extralymphoid Effector Memory Cell Population in Humans1
http://www.100md.com
免疫学杂志 2005年第5期
Abstract
Although mouse studies have demonstrated the presence of an effector memory population in nonlymphoid tissues, the phenotype of human CD8+ T cells present in such compartments has not been characterized. Because of the relatively large number of CD8+ T cells present in breast milk, we were able to characterize the phenotype of this cell population in HIV-infected and uninfected lactating women. CMV, influenza virus, EBV, and HIV-specific CD8+ T cells as measured by the IFN- ELISPOT and MHC class I tetramer staining were all present at greater frequencies in breast milk as compared with blood. Furthermore, a greater percentage of the breast milk CD8+ T cells expressed the intestinal homing receptor, CD103, and the mucosal homing receptor CCR9. Breast milk T cells were predominantly CD45RO+HLADR+ and expressed low levels of CD45RA, CD62L, and CCR7 consistent with an effector memory population. Conversely, T cells derived from blood were mainly characterized as central memory cells (CCR7+CD62L+). These results demonstrate a population of extralymphoid CD8+ T cells with an effector memory phenotype in humans, which could contribute to enhanced local virologic control and the relative lack of HIV transmission via this route.
Introduction
Immunologic memory provides long-term protection against infection due to the presence of subsets of T cells that are present long after initial Ag encounter. These memory T cells survive beyond the acute period of infection when the majority of cells expand and function to control the initial infection then apoptose (1). These remaining memory T cells are capable of responding rapidly to pathogen re-encounter because they are more numerous, are capable of persisting in the absence of Ag, and proliferate much more rapidly when compared with naive T cells (2). These T cells are also heterogeneous in their expression of surface molecules, effector function, and distribution. They have been defined by several different parameters including the lymph node homing receptors CCR7 and CD62L (3), costimulatory molecules CD27 and CD28 (4, 5, 6), and activation state using CD45RA and CD45RO (4, 7).
These types of analyses have led to the description of two main subsets of memory T cells: central memory T cells characterized by expression of CCR7 and CD62L and residing in secondary lymphoid organs, and effector memory T cells lacking expression of CCR7 and CD62L with localization in nonlymphoid compartments. Effector memory T cells were found to acquire effector functions, such as IFN- secretion and killing capabilities more quickly than central memory T cells (3, 8, 9). Moreover, these Ag-specific CD8 T cells were shown to migrate to various organs, and recent data suggested the existence of an extralymphoid effector memory T cell population capable of immediate response against pathogens in mouse models (9). These studies also showed that CD8 effector memory cells preferentially migrated and expanded in nonlymphoid tissue and that the number of Ag-specific cells within the nonlymphoid compartment represented a substantial portion of the overall response. As would be expected, these cells lacked the lymph node homing receptors CCR7 and CD62L (3, 9, 10). Studies of central and effector memory T cells in humans are hampered by difficulties in obtaining peripheral nonlymphoid tissues. Furthermore, even when these tissues are available for study, the number of Ag-specific T cells obtained is too small to adequately evaluate. Recently, we and others (11, 12) demonstrated the presence of Ag-specific CD8+ T cells in breast milk of HIV-positive women. These cells were present in relatively large numbers and were readily available from women agreeing to donate breast milk postpartum. We took advantage of these characteristics and used the cells derived from breast milk as a source of CD8+ T cells from nonlymphoid sites.
The current study demonstrates that the magnitude and quality of T cells in breast milk are different when compared with the peripheral blood compartment. Additionally, we find that these nonlymphoid-derived CD8+ T cells are more activated and display markers indicative of an effector memory T cell population relative to the central memory phenotype displayed by PBMC. Breast milk cells (BMC) also express the mucosal homing receptors CD103 and CCR9 suggesting that they may have originated in the gastrointestinal tract. Last, these data demonstrate that this population of nonlymphoid cells, previously characterized in mice (9), is present in humans as well.
Materials and Methods
Subjects and HLA typing
Lactating HIV-negative (n = 11) and HIV-positive (n = 7) women were recruited from University of Alabama at Birmingham (UAB) to donate breast milk and blood. Written informed consent was obtained from all women who participated in this study. The Institutional Review Board of the University of Alabama approved the UAB study. HLA typing was performed at the Tissue Typing Center in the UAB Hospital using the Micro SSP HLA typing system (One Lamba).
Isolation of breast milk cells
Breast milk expressed using a breast pump was obtained from women 2 days to 4.5 mo postpartum. Milk samples were kept at 4°C for no >5 h and then milk was centrifuged at 400 x g for 15 min. Cream that formed at the top was scooped out with a sterile spatula, milk supernatant was saved for viral load determinations, and the cell pellet was washed two times with HBSS (12). Cells were resuspended in complete RPMI 1640 with 10% AB sera and viable BMC were counted using a hemocytometer by trypan blue exclusion. PBMC were obtained by standard Histopaque density centrifugation (Sigma-Aldrich).
IFN- ELISPOT assay
An IFN- ELISPOT assay was used to enumerate HIV-specific T cells from breast milk or PBMC of HIV-infected or uninfected women (12). Briefly, 96-well nitrocellulose plates (Milliliter HA; Millipore) were coated with 5 μg/ml mouse anti-human IFN- mAb (clone 1-D1K; Mabtech) and then incubated overnight at 4°C. Ab was decanted, and wells were blocked with 200 μl of RPMI 1640 containing 10% human AB serum. BMC were added to the plates at a concentration of 5 x 105 cells/ml and incubated overnight with pools of 20-mer peptides (overlapping by 10 aa) spanning the entire sequence of HIV-1 clade B Gag (HXB2), Pol (HXB2), Env (MN), or Nef (BRU) (AIDS Research and Reference Reagent Program, Division of AIDS, National Institute of Allergy and Infectious Diseases (NIAID), National Institutes of Health) at 2 μg/ml. After washing, 1 μg/ml biotinylated anti-IFN- mAb 7-B6-1 (Mabtech) was added to the plates for 2 h at room temperature. Following another round of washing, streptavidin-conjugated alkaline phosphatase (Southern Biotechnologies) was added, and the plates were incubated at room temperature for an additional hour. Last, NBT/5-bromo-4-chloro-3-indolyl phosphate; Sigma-Aldrich) was added for color development. Individual cytokine-producing cells were counted manually and/or by the C.T.L. ImmunoSpot analyzer (Cellular Technologies). In addition, immune responses to HLA-restricted peptides (8–10 mers) from HIV-1, CMV, EBV, and influenza were also measured using the ELISPOT at a concentration of 10–5 M (13). PHA (Sigma-Aldrich) (5 μg/ml), was used as a positive control and unstimulated cells, i.e., medium alone, as the negative control. Responses were considered positive if there were 100 spot-forming cells (SFC)4/106 cells and twice as many SFC as in the nonstimulated control wells. PBMC were added to the ELISPOT plates at a concentration of 1 x 106 cells/ml and incubated with the same peptide pools as the BMC. The SFC was normalized to SFC per CD8+ T cells by using the percentage of CD8+ cells present in either breast milk or PBMC using flow cytometry.
Quantitation of HIV-1 RNA in plasma and breast milk
HIV RNA levels in the plasma and breast milk samples were determined by the Amplicor UltraSensitive HIV-1 Monitor version 1.5 assay (Roche Diagnostics Systems), according to the manufacturer’s protocol. We have previously demonstrated that this assay can reproducibly and reliably detect HIV in breast milk (14). In addition, we and others (15, 16, 17) have shown that breast milk RNA levels are at least 10- to 100-fold less than those found in the blood. The breast milk RNA levels in the HIV-infected women in this study were all <50 copies/ml.
Surface marker staining
Phenotypic characterization of BMC and PBMC used cell-surface markers CD3, CD4, CD8, CD57, CD62L, CD103, HLA-DR, CD45RO, CD45RA, CD27, CD28, CD38, CCR7, and CCR9 (BD Biosciences). Stained cells were acquired using a BD Calibur flow cytometer (BD Biosciences) and analyzed with FlowJo version 4.3.1 software (Tree Star). Our design was to run 50,000 gated T cells (based on CD3 staining) for each stained specimen. When CD3 was not used as a marker, we gated on CD8-staining cells.
MHC class I tetramer analysis
Enumeration of epitope-specific cells by MHC class I tetramer complexes was performed using either HLA B7 CMV pp65 TPRVTGGGAM, HLA A2 HIV Gag p17 SLYNTVATL, HLA A2 HIV Pol ILKEPVHGV, HLA A3 HIV Gag p17 RLRPGGKKK, or HLA B35 HIV Pol NPDDIVIYQY (kindly provided by L. Tussey, Merck, West Point, PA) tetramers. All HIV-specific MHC class I tetramers were synthesized by the NIAID Tetramer Facility (AIDS Research and Reference Reagent Program, Division of AIDS, NIAID, National Institutes of Health) and conjugated with PE. B7-CMV was kindly provided by S. Boppana (University of Alabama at Birmingham) and was conjugated to allophycocyanin. Tetramer titration and staining conditions were maximized by using epitope-specific PBMC from appropriate HLA seropositive and seronegative volunteers. For each analysis, a minimum of 10,000 events are represented.
Statistics
Comparison of the frequency and phenotype of T cells between breast milk and blood compartments were made by the nonparametric Mann-Whitney U test using Analyze-It software.
Results
Study population
BM cells and PBMC from 18 lactating women were analyzed for phenotypic differences, seven were HIV positive and 11 were HIV negative. Breast milk was collected between 2 days and 4.5 mo postpartum with most samples being collected within 1 wk of delivery. Median plasma viral loads for the HIV-positive women were <50 copies/ml, and the median CD4 count was 387 cells/mm3. All HIV-positive women were on antiretroviral therapy.
Comparison of the frequency of Ag-specific CD8+ T cells in breast milk and blood
Ag-specific IFN--secreting CD8+ T cell responses measured in breast milk and blood were normalized to CD8 SFC/105 T cells so that direct comparisons could be made. Most responses found in breast milk were significantly increased when compared with PBMC. These findings applied to CD8+ T cells stimulated with HIV-specific peptide pools (Fig. 1, A–C) as well as to those stimulated with HLA class I-restricted peptides from HIV, EBV, CMV, and influenza and matched for the particular individuals HLA class I (Fig. 1, D–F).
FIGURE 1. HIV- and non-HIV-specific immune responses as measured by the IFN- ELISPOT assay normalized to the number of CD8+ T cells. A–C, PBMC and BMC were stimulated with overlapping 20-mer peptide pools from Gag, Pol, Env, and Nef. D–F, Immune responses to HLA-restricted peptides (8–10 mers) from HIV, CMV, EBV, and influenza. A2-Flu (GILGFVFTL); A3-Flu (ILRGSVAHK); A3-EBV EBNA3A (RLRAEAQVK); A3-EBV BRLF1 (RVRAYTYSK); A11-EBV EBNA3B (IVTDFSVIK); A11-EBV BRLF1 (ATIGTAMYK); B7-CMV (TPRVTGGGAM), B8-EBV EBNA3A (FLRGRAYGL). Representative BMC and PBMC are taken from HIV-seropositive women. Note that the responses are plotted on a log scale since some of the differences were large. Responses were measured for both breast milk and blood for all Ags shown; thus, lack of a response signifies that response was not detected.
Next, we performed HLA class I tetramer staining in three women who had detectable responses to HIV and/or CMV. All displayed increased frequency of tetramer binding T cells in breast milk when compared with PBMC (Table I). This is consistent with our prior studies wherein we found that the tetramer frequency in blood was only 0.22% compared with 0.65% in breast milk (12).
Table I. Class I tetramer staining comparison between BMC and PBMC
The data for the B7-CMV tetramer was derived from the same volunteer used to detect B7-CMV-specific peptide responses in the ELISPOT assay (Fig. 1F). Of note, although increased responses are seen for breast milk cells in this volunteer with both IFN- ELISPOT assay (Fig. 1F, 2/3 non-HIV Ags) and tetramer (Table I), the differences are not as pronounced as those seen for the other volunteers.
Intracellular IFN- staining after PBMC stimulation obtained from another HIV-infected woman with the A2-SL9 peptide also demonstrated a higher frequency of IFN--secreting CD8+ T cells in breast milk (1.72%) when compared with PBMC (0.13%) (data not shown). Thus, with few exceptions, the vast majority of responses were detected at higher frequencies in breast milk.
Comparison of surface marker expression between BMC and PBMC
Given the differences in frequency of CD8+ Ag-specific T cells noted between BMC and PBMC, we were interested in determining whether the phenotype of these two populations of lymphocytes differed, especially since they represented lymphocytes from lymphoid and nonlymphoid compartments. When looking at standard lymphocyte surface markers, we noted that the median CD3+/CD8+ BMC percentages in both HIV positive and HIV negative were slightly greater when compared with PBMC (Table II). However, the total number of lymphocytes per milliliter present in breast milk is lower than PBMC (data not shown). The only significant difference when comparing HIV-positive to HIV-negative women was the finding of consistently lower percentage of CD4 T cells in the former group in PBMC (29.4% vs 54%, p = 0.012), and lower CD103 in BMC of HIV-positive women when compared with HIV negative (14.2% vs 32.2%, p = 0.003) (Table II). All other surface cell phenotypic markers were not statistically different when comparing HIV-positive to HIV-negative women.
Table II. Surface marker expression comparison between BMC and PBMC
Expression of CD103, an integrin directed to E-cadherin was at least an order of magnitude higher in BMC compared with PBMC in both HIV-positive and HIV-negative women (Table II and Fig. 2). Increased CD103 is suggestive of BMC CD8+ cell migration from epithelial compartments of the gastrointestinal tract. CCR9, another mucosal homing receptor, was also significantly higher in BMC of HIV-seronegative women when compared with their PBMC (Table II and Fig. 2).
FIGURE 2. Surface marker expression on CD8+ T cells from 18 lactating women. A, HIV-seropositive women (n = 7); B, HIV-seronegative women (n = 11). Surface marker expression on BMC vs PBMC is shown. Cells were gated by using the following parameters: forward scatter vs side scatter and CD3+/CD8+ or CD8+ T cells if CD3 was not performed. The median percentage of CD8+ T cells that stained with the given surface marker is plotted.
Reduced levels of CCR7 expression were detected on BMC (0.8% in HIV positive and 2.1% in HIV negative), whereas PBMC contained levels of expression as seen in other studies (28.2% in HIV positive and 52.2% in HIV negative) (10, 18). Furthermore, the large majority of T cells in breast milk from either HIV-seropositve or -seronegative women had a memory phenotype, as they expressed high levels of CD45RO, and reduced expression of CD45RA and the lymph node homing marker CD62L. The activation marker, HLA-DR, was highly expressed on the surface of breast milk-derived CD8+ T cells relative to PBMC in both HIV-positive and HIV-negative women (Fig. 2 and Table II). In contrast, breast milk from both HIV-positive and HIV-negative women and PBMC from HIV-positive women expressed high levels of another activation marker CD38. Levels of CD38 in PBMC of HIV-negative women were lower but not significantly different (Table II). Similar levels of CD57 expression, a marker suggestive of late memory T cells with little proliferative capacity (19, 20, 21) were observed when comparing the level of expression between BMC and PBMC from either HIV-seropositive or HIV-seronegative women (Table II and Fig. 2).
In short, the measured surface phenotypes on most T cell populations in the BMC samples from HIV-seronegative women differed significantly from the populations in the corresponding peripheral blood, with respect to all tested T cell markers except for CD38, CD57, CD4, and CD8 (Table II). The same trends were noted for BMC derived from HIV-positive women, but statistically significant differences were not as consistently observed most likely due to a small sample size.
Comparison of surface marker expression on Ag-specific CD8+ T cells from BMC and PBMC
The above data showed marked differences in the surface marker expression on whole populations of BMC vs PBMC. We next evaluated whether these differences were present in Ag-specific populations. Cells from two HIV-infected lactating women with known tetramer-positive responses were studied. Responses to B35Pol-NY9 were from volunteer 1 and responses to A2Pol-IV9 and B7-CMV from volunteer 2 (Table I). Similar to the finding with the whole T cell populations, differences were observed in the expression of CD45RA (lower in BMC from both HIV- and CMV-specific populations). HLA-DR also showed greater expression on the surface of BMC than PBMC. These differences occurred whether the Ag-specific cells were HIV or CMV specific (Fig. 3). We also determined the relative differentiation status of these Ag-specific cells by using the expression of the co-stimulatory receptors CD28 and CD27. These receptors are involved in the regulation of T cell activation and in the generation of Ag-primed cells, respectively, and have been used by others to distinguish between subsets of differentiated CD8+ T cells (4, 6). CD28 seems to be expressed at higher levels on the surface of BMC, whereas CD27 appears to be present at similar levels on both BMC and PBMC (Fig. 3).
FIGURE 3. Surface marker expression on Ag-specific cells from two HIV-infected women. Cells were gated on CD8+ tetramer + cells. HIV-1 Pol (aa 330–338, NPDIVIYQY) HLA B35 restricted staining, HIV-1 Pol (aa 309–317, ILKEPVHGV) HLA A2 restricted staining and CMV pp65 (aa 417–426, TPRVTGGGAM) HLA B7
Discussion
Memory T cells have been divided into multiple functional phenotypes on the basis of surface molecule expression, anatomic location, and immune function. Currently, there is no clear consensus on the pathways of memory/effector T cell differentiation, and several models have been proposed (2, 3, 6, 7, 22, 23, 24, 25, 26, 27). Studies in mice suggest that nonlymphoid tissues are preferentially populated by a subset of CD8+ T cells (9, 28). These cells express low levels of CCR7 and CD62L and are capable of producing effector cytokines immediately upon Ag stimulation; they have been referred to as "effector" memory T cells (Tem). In the present study, we characterized the phenotype of CD8+ T cells derived from breast milk from both HIV-infected and uninfected women. Overall, the phenotype of breast milk cells in these populations were remarkably similar, no doubt reflecting the fact all the HIV-infected women had normal levels of CD4 cells and were on highly active antiretroviral therapy. Using both ELISPOT and tetramer assays, we found that the frequency of Ag-specific CD8+ T cells in breast milk is significantly higher than PBMC. This is consistent with our previously published work as well as that of others (11, 12). Higher levels of Ag could not explain the higher frequency of Ag-specific cells, since in the case of HIV infection, breast milk contains much less HIV than plasma (12, 17, 29). In addition, compared with their counterparts in blood, breast milk CD8+ T cells lack expression of CCR7, CD62L, and CD45RA and express high levels of CD45RO. Breast milk T cells also express higher levels of the activation marker HLA-DR. Thus, the lactating breast is enriched for Ag-specific T cells distinct from those circulating in the blood, and these cells bear phenotypic hallmarks of effector memory cells that have been recently activated.
Expression of the co-stimulatory molecules CD27 and CD28 have also been associated with different stages of T cell differentiation (4, 6, 30). Sequential down-regulation of CD28 and then CD27 expression has been described during CD8+ T cell differentiation (4, 31). Our data on the expression of these markers for CMV-specific responses in PBMC of a lactating HIV-positive woman are consistent with that described by others (4, 27). CMV-specific cells express levels of CD27 and CD28 similar to those reported (4, 6), with concomitant expression of CD45RA showing as others have that these cells are in an intermediate to late stage of differentiation (4, 27). The low expression of CD45RA and high expression of HLA-DR on all Ag-specific breast milk cells (Fig. 3) is consistent with our data on whole BMC populations (Fig. 2 and Table II). Although we did not measure CCR7 on Ag-specific cells, it is unlikely that these cells would express this marker since very few of the total breast milk-derived population of CD8+
T cells expressed CCR7 (0.8% in HIV positive and 2.1% in HIV negative) (Table II and Fig. 2). Moreover, studies of Ag-specific T cells have demonstrated that greater than 70% of the Ag-specific CD8+ T cells do not express CCR7 (4, 7).
Recent studies have defined two subsets of effector memory (CD45RA–CCR7–) T cell subsets in humans, one expressing CD27 (CD27+) and one that does not (CD27–) (32). In our Ag-specific breast milk-derived CD8+ T cell populations, we have both of these populations represented (Fig. 3). In humans, the CD45RA–CD27+ subset is capable of cytolytic activity after in vitro stimulation (31). We have previously demonstrated that CD8+ breast milk-derived T cells were also capable of Ag-specific cytolysis after in vitro stimulation (12).
The expression of CD28 on breast milk Ag-specific cells was relatively high. According to some models of phenotypic evolution, expression of this marker is a sign of "earlier" differentiation and suggests that breast milk cells may be at an earlier stage in postthymic development than those cells found in the blood. Presumably this is due to less Ag in breast milk and/or exposure to other factors in the unique mucosal environment of the lactating breast. In addition, this may have something to do with the recent acquisition/homing of these cells into this compartment, given that breast milk is not produced before pregnancy. Given the limited number of Ag-specific samples analyzed, it is difficult to make definitive conclusions, and analysis of more samples is needed.
The origin of breast milk T cells is uncertain. Animal studies clearly demonstrate that B cells from Peyer’s patches migrate specifically to the lactating mammary gland (33, 34, 35, 36) and that pregnancy increases the mucosal vascular addressin MadCAM-1, which interacts with the gut homing receptor 47 (37). In humans, oral immunization with nonpathogenic Escherichia coli, produces type-specific IgA in breast milk but not in serum or saliva of lactating women, suggesting trafficking between breast and blood (38). There are few data on T cell migration between the intestine and the lactating breast. Labeling studies in rats and pigs demonstrate that T cells in the breast originate from the gut as well as blood (39, 40). We studied the expression of two mucosal markers CD103 (E7) and CCR9 in breast milk lymphocytes. CD103 is expressed on intraepithelial lymphocytes in the gut, genitourinary tract, and skin, as well as its increased expression in activated CD8+ cells (41). In contrast to PBMC where CD103 is expressed on very few lymphocytes (<2%), we found that it was expressed on a significant number of breast milk lymphocytes (32.2% in HIV negative and 14.2% in HIV positive), suggesting selective recruitment or retention of these cells. CCR9 is a chemokine receptor expressed on thymocytes, intestinal lymphocytes, and on discrete subsets of T cells that traffic to the intestine (42, 43). Expression of CCR9 on breast milk cells was significantly greater than that found in blood in HIV-negative women. Thus, expression of these mucosal homing receptors by CD8+ T cells from breast milk suggests that many of these cells may have originated from the gastrointestinal tract.
Taken together, these data demonstrate that breast milk is populated with CD8+ T cells that have an effector memory phenotype, consistent with studies in animals showing that a substantial portion of the T cell response to infection is focused on nonlymphoid tissue (1, 9, 44, 45, 46, 47). Study of breast milk cells provides a novel convenient method for the study of these cells in humans. Given the importance of milk in protecting progeny from infection, the enrichment of Ag-specific cells with an effector phenotype is teleologically appealing. The immunologic benefits of breast-feeding are well established (48), but the role of these effector memory T cells in the prevention of infection remains to be elucidated.
Disclosures
The authors have no financial conflict of interest.
Acknowledgments
We thank the volunteers for their participation, Barbara Corley and Shawn Dillon for breast milk procurement, Marion Spell and Tracy McGuire for help with the flow cytometric acquisition, Anju Bansal for insightful discussions, Linda Tussey for providing the B35 tetramer, Suresh Boppana for providing the B7 CMV tetramer and the Children’s Hospital Family Clinic.
Footnotes
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
1 This work was supported by grants from the National Institutes of Health (R01 HD-40777, HD-396110, AI-049126) and the Elizabeth Glaser Pediatric AIDS Foundation. G.M.A. is an Elizabeth Glaser Pediatric AIDS Foundation Scientist.
2 S.S. and M.K.G. contributed equally to this work.
3 Address correspondence and reprint requests to Dr. Grace M. Aldrovandi, Infectious Diseases #51, Children’s Hospital Los Angeles, 4650 Sunset Boulevard, Los Angeles, CA 90027. E-mail address: galdrovandi{at}chla.usc.edu
4 Abbreviation used in this paper: SFC, spot-forming cells.
Received for publication July 19, 2004. Accepted for publication December 20, 2004.
References
Masopust, D., V. Vezys, E. J. Usherwood, L. S. Cauley, S. Olson, A. L. Marzo, R. L. Ward, D. L. Woodland, L. Lefrancois. 2004. Activated primary and memory CD8 T cells migrate to nonlymphoid tissues regardless of site of activation or tissue of origin. J. Immunol. 172:4875.
Wherry, E. J., V. Teichgraber, T. C. Becker, D. Masopust, S. M. Kaech, R. Antia, U. H. von Andrian, R. Ahmed. 2003. Lineage relationship and protective immunity of memory CD8 T cell subsets. Nat. Immunol. 4:225.
Sallusto, F., D. Lenig, R. Forster, M. Lipp, A. Lanzavecchia. 1999. Two subsets of memory T lymphocytes with distinct homing potentials and effector functions. Nature 401:708.
Appay, V., P. R. Dunbar, M. Callan, P. Klenerman, G. M. Gillespie, L. Papagno, G. S. Ogg, A. King, F. Lechner, C. A. Spina, et al 2002. Memory CD8+ T cells vary in differentiation phenotype in different persistent virus infections. Nat. Med. 8:379.
Tomiyama, H., T. Matsuda, M. Takiguchi. 2002. Differentiation of human CD8+ T cells from a memory to memory/effector phenotype. J. Immunol. 168:5538.
Tussey, L. G., U. S. Nair, M. Bachinsky, B. H. Edwards, J. Bakari, K. Grimm, J. Joyce, R. Vessey, R. Steigbigel, M. N. Robertson, et al 2003. Antigen burden is major determinant of human immunodeficiency virus-specific CD8+ T cell maturation state: potential implications for therapeutic immunization. J. Infect. Dis. 187:364.
Champagne, P., G. S. Ogg, A. S. King, C. Knabenhans, K. Ellefsen, M. Nobile, V. Appay, G. P. Rizzardi, S. Fleury, M. Lipp, et al 2001. Skewed maturation of memory HIV-specific CD8 T lymphocytes. Nature 410:106.
Sabbaj, S., B. H. Edwards, M. K. Ghosh, K. Semrau, S. Cheelo, D. M. Thea, L. Kuhn, G. D. Ritter, M. J. Mulligan, P. A. Goepfert, G. M. Aldrovandi. 2002. Human immunodeficiency virus-specific CD8+ T cells in human breast milk. J. Virol. 76:7365.
Sabbaj, S., A. Bansal, G. D. Ritter, C. Perkins, B. H. Edwards, E. Gough, J. Tang, J. J. Szinger, B. Korber, C. M. Wilson, et al 2003. Cross-reactive CD8+ T cell epitopes identified in US adolescent minorities. J. Acquir. Immune Defic. Syndr. 33:426
Ghosh, M. K., L. Kuhn, J. West, K. Semrau, D. Decker, D. M. Thea, G. M. Aldrovandi. 2003. Quantitation of human immunodeficiency virus type 1 in breast milk. J. Clin. Microbiol. 41:2465.
Nduati, R., G. John. 1995. Breast milk transmission of HIV-1. NARESA Mongr Dec:1.
Semba, R. D., N. Kumwenda, D. R. Hoover, T. E. Taha, T. C. Quinn, L. Mtimavalye, R. J. Biggar, R. Broadhead, P. G. Miotti, L. J. Sokoll, et al 1999. Human immunodeficiency virus load in breast milk, mastitis, and mother-to-child transmission of human immunodeficiency virus type 1. J. Infect. Dis. 180:93.
Willumsen, J. F., M. L. Newell, S. M. Filteau, A. Coutsoudis, S. Dwarika, D. York, A. M. Tomkins, H. M. Coovadia. 2001. Variation in breastmilk HIV-1 viral load in left and right breasts during the first 3 months of lactation. AIDS 15:1896.
Chen, G., P. Shankar, C. Lange, H. Valdez, P. R. Skolnik, L. Wu, N. Manjunath, J. Lieberman. 2001. CD8 T cells specific for human immunodeficiency virus, Epstein-Barr virus, and cytomegalovirus lack molecules for homing to lymphoid sites of infection. Blood 98:156.
Brenchley, J. M., N. J. Karandikar, M. R. Betts, D. R. Ambrozak, B. J. Hill, L. E. Crotty, J. P. Casazza, J. Kuruppu, S. A. Migueles, M. Connors, et al 2003. Expression of CD57 defines replicative senescence and antigen-induced apoptotic death of CD8+ T cells. Blood 101:2711.
d’Angeac, A. D., S. Monier, D. Pilling, A. Travaglio-Encinoza, T. Reme, M. Salmon. 1994. CD57+ T lymphocytes are derived from CD57– precursors by differentiation occurring in late immune responses. Eur. J. Immunol. 24:1503
Kern, F., E. Khatamzas, I. Surel, C. Frommel, P. Reinke, S. L. Waldrop, L. J. Picker, H. D. Volk. 1999. Distribution of human CMV-specific memory T cells among the CD8pos. subsets defined by CD57, CD27, and CD45 isoforms. Eur. J. Immunol. 29:2908
Ahmed, R., D. Gray. 1996. Immunological memory and protective immunity: understanding their relation. Science 272:54.
Kaech, S. M., E. J. Wherry, R. Ahmed. 2002. Effector and memory T-cell differentiation: implications for vaccine development. Nat. Rev. Immunol. 2:251.
Kaech, S. M., J. T. Tan, E. J. Wherry, B. T. Konieczny, C. D. Surh, R. Ahmed. 2003. Selective expression of the interleukin 7 receptor identifies effector CD8 T cells that give rise to long-lived memory cells. Nat. Immunol. 4:1191.
Seder, R. A., R. Ahmed. 2003. Similarities and differences in CD4+ and CD8+ effector and memory T cell generation. Nat. Immunol. 4:835.
Lanzavecchia, A., F. Sallusto. 2002. Progressive differentiation and selection of the fittest in the immune response. Nat. Rev. Immunol. 2:982.
Appay, V., S. L. Rowland-Jones. 2004. Lessons from the study of T-cell differentiation in persistent human virus infection. Semin. Immunol. 16:205
Reinhardt, R. L., A. Khoruts, R. Merica, T. Zell, M. K. Jenkins. 2001. Visualizing the generation of memory CD4 T cells in the whole body. Nature 410:101.
Rousseau, C. M., R. W. Nduati, B. A. Richardson, M. S. Steele, G. C. John-Stewart, D. A. Mbori-Ngacha, J. K. Kreiss, J. Overbaugh. 2003. Longitudinal analysis of human immunodeficiency virus type 1 RNA in breast milk and of its relationship to infant infection and maternal disease. J. Infect. Dis. 187:741.
Hamann, D., M. T. Roos, R. A. van Lier. 1999. Faces and phases of human CD8 T-cell development. Immunol. Today 20:177.
Hamann, D., P. A. Baars, M. H. Rep, B. Hooibrink, S. R. Kerkhof-Garde, M. R. Klein, R. A. van Lier. 1997. Phenotypic and functional separation of memory and effector human CD8+ T cells. J. Exp. Med. 186:1407.
Rufer, N., A. Zippelius, P. Batard, M. J. Pittet, I. Kurth, P. Corthesy, J. C. Cerottini, S. Leyvraz, E. Roosnek, M. Nabholz, P. Romero. 2003. Ex vivo characterization of human CD8+ T subsets with distinct replicative history and partial effector functions. Blood 102:1779.
Roux, M. E., M. McWilliams, J. M. Phillips-Quagliata, M. E. Lamm. 1981. Differentiation pathway of Peyer’s patch precursors of IgA plasma cells in the secretory immune system. Cell. Immunol. 61:141
Tanneau, G. M., L. Hibrand-Saint Oyant, C. C. Chevaleyre, H. P. Salmon. 1999. Differential recruitment of T- and IgA B-lymphocytes in the developing mammary gland in relation to homing receptors and vascular addressins. J. Histochem. Cytochem. 47:1581
Goldblum, R. M., S. Ahlstedt, B. Carlsson, L. A. Hanson, U. Jodal, G. Lidin-Janson, A. Sohl-Akerlund. 1975. Antibody-forming cells in human colostrum after oral immunisation. Nature 257:797.
Manning, L. S., M. J. Parmely. 1980. Cellular determinants of mammary cell-mediated immunity in the rat. I. The migration of radioisotopically labeled T lymphocytes. J. Immunol. 125:2508.
Salmon, H.. 2000. Mammary gland immunology and neonate protection in pigs. Homing of lymphocytes into the MG. Adv. Exp. Med. Biol. 480:279.
Agace, W. W., J. M. Higgins, B. Sadasivan, M. B. Brenner, C. M. Parker. 2000. T-lymphocyte-epithelial-cell interactions: integrin (E)(CD103)(7), LEEP-CAM and chemokines. Curr. Opin. Cell Biol. 12:563.
Zabel, B. A., W. W. Agace, J. J. Campbell, H. M. Heath, D. Parent, A. I. Roberts, E. C. Ebert, N. Kassam, S. Qin, M. Zovko, et al 1999. Human G protein-coupled receptor GPR-9-6/CC chemokine receptor 9 is selectively expressed on intestinal homing T lymphocytes, mucosal lymphocytes, and thymocytes and is required for thymus-expressed chemokine-mediated chemotaxis. J. Exp. Med. 190:1241.
Svensson, M., J. Marsal, A. Ericsson, L. Carramolino, T. Broden, G. Marquez, W. W. Agace. 2002. CCL25 mediates the localization of recently activated CD8+ lymphocytes to the small-intestinal mucosa. J. Clin. Invest. 110:1113.
Klonowski, K. D., K. J. Williams, A. L. Marzo, D. A. Blair, E. G. Lingenheld, L. Lefrancois. 2004. Dynamics of blood-borne CD8 memory T cell migration in vivo. Immunity 20:551
La Gruta, N. L., S. J. Turner, P. C. Doherty. 2004. Hierarchies in cytokine expression profiles for acute and resolving influenza virus-specific CD8+ T cell responses: correlation of cytokine profile and TCR avidity. J. Immunol. 172:5553.
Sun, J. C., M. J. Bevan. 2004. Cutting edge: long-lived CD8 memory and protective immunity in the absence of CD40 expression on CD8 T cells. J. Immunol. 172:3385.
Wherry, E. J., J. N. Blattman, K. Murali-Krishna, R. van der Most, R. Ahmed. 2003. Viral persistence alters CD8 T-cell immunodominance and tissue distribution and results in distinct stages of functional impairment. J. Virol. 77:4911
Fituch, C., K. Palkowetz, A. Goldman, R. Schanler. 2004. Concentrations of IL-10 in preterm human milk and in milk from mothers of infants with necrotizing enterocolitis. Acta Paediatr. 93:1496(Steffanie Sabbaj, Mrinal )
Although mouse studies have demonstrated the presence of an effector memory population in nonlymphoid tissues, the phenotype of human CD8+ T cells present in such compartments has not been characterized. Because of the relatively large number of CD8+ T cells present in breast milk, we were able to characterize the phenotype of this cell population in HIV-infected and uninfected lactating women. CMV, influenza virus, EBV, and HIV-specific CD8+ T cells as measured by the IFN- ELISPOT and MHC class I tetramer staining were all present at greater frequencies in breast milk as compared with blood. Furthermore, a greater percentage of the breast milk CD8+ T cells expressed the intestinal homing receptor, CD103, and the mucosal homing receptor CCR9. Breast milk T cells were predominantly CD45RO+HLADR+ and expressed low levels of CD45RA, CD62L, and CCR7 consistent with an effector memory population. Conversely, T cells derived from blood were mainly characterized as central memory cells (CCR7+CD62L+). These results demonstrate a population of extralymphoid CD8+ T cells with an effector memory phenotype in humans, which could contribute to enhanced local virologic control and the relative lack of HIV transmission via this route.
Introduction
Immunologic memory provides long-term protection against infection due to the presence of subsets of T cells that are present long after initial Ag encounter. These memory T cells survive beyond the acute period of infection when the majority of cells expand and function to control the initial infection then apoptose (1). These remaining memory T cells are capable of responding rapidly to pathogen re-encounter because they are more numerous, are capable of persisting in the absence of Ag, and proliferate much more rapidly when compared with naive T cells (2). These T cells are also heterogeneous in their expression of surface molecules, effector function, and distribution. They have been defined by several different parameters including the lymph node homing receptors CCR7 and CD62L (3), costimulatory molecules CD27 and CD28 (4, 5, 6), and activation state using CD45RA and CD45RO (4, 7).
These types of analyses have led to the description of two main subsets of memory T cells: central memory T cells characterized by expression of CCR7 and CD62L and residing in secondary lymphoid organs, and effector memory T cells lacking expression of CCR7 and CD62L with localization in nonlymphoid compartments. Effector memory T cells were found to acquire effector functions, such as IFN- secretion and killing capabilities more quickly than central memory T cells (3, 8, 9). Moreover, these Ag-specific CD8 T cells were shown to migrate to various organs, and recent data suggested the existence of an extralymphoid effector memory T cell population capable of immediate response against pathogens in mouse models (9). These studies also showed that CD8 effector memory cells preferentially migrated and expanded in nonlymphoid tissue and that the number of Ag-specific cells within the nonlymphoid compartment represented a substantial portion of the overall response. As would be expected, these cells lacked the lymph node homing receptors CCR7 and CD62L (3, 9, 10). Studies of central and effector memory T cells in humans are hampered by difficulties in obtaining peripheral nonlymphoid tissues. Furthermore, even when these tissues are available for study, the number of Ag-specific T cells obtained is too small to adequately evaluate. Recently, we and others (11, 12) demonstrated the presence of Ag-specific CD8+ T cells in breast milk of HIV-positive women. These cells were present in relatively large numbers and were readily available from women agreeing to donate breast milk postpartum. We took advantage of these characteristics and used the cells derived from breast milk as a source of CD8+ T cells from nonlymphoid sites.
The current study demonstrates that the magnitude and quality of T cells in breast milk are different when compared with the peripheral blood compartment. Additionally, we find that these nonlymphoid-derived CD8+ T cells are more activated and display markers indicative of an effector memory T cell population relative to the central memory phenotype displayed by PBMC. Breast milk cells (BMC) also express the mucosal homing receptors CD103 and CCR9 suggesting that they may have originated in the gastrointestinal tract. Last, these data demonstrate that this population of nonlymphoid cells, previously characterized in mice (9), is present in humans as well.
Materials and Methods
Subjects and HLA typing
Lactating HIV-negative (n = 11) and HIV-positive (n = 7) women were recruited from University of Alabama at Birmingham (UAB) to donate breast milk and blood. Written informed consent was obtained from all women who participated in this study. The Institutional Review Board of the University of Alabama approved the UAB study. HLA typing was performed at the Tissue Typing Center in the UAB Hospital using the Micro SSP HLA typing system (One Lamba).
Isolation of breast milk cells
Breast milk expressed using a breast pump was obtained from women 2 days to 4.5 mo postpartum. Milk samples were kept at 4°C for no >5 h and then milk was centrifuged at 400 x g for 15 min. Cream that formed at the top was scooped out with a sterile spatula, milk supernatant was saved for viral load determinations, and the cell pellet was washed two times with HBSS (12). Cells were resuspended in complete RPMI 1640 with 10% AB sera and viable BMC were counted using a hemocytometer by trypan blue exclusion. PBMC were obtained by standard Histopaque density centrifugation (Sigma-Aldrich).
IFN- ELISPOT assay
An IFN- ELISPOT assay was used to enumerate HIV-specific T cells from breast milk or PBMC of HIV-infected or uninfected women (12). Briefly, 96-well nitrocellulose plates (Milliliter HA; Millipore) were coated with 5 μg/ml mouse anti-human IFN- mAb (clone 1-D1K; Mabtech) and then incubated overnight at 4°C. Ab was decanted, and wells were blocked with 200 μl of RPMI 1640 containing 10% human AB serum. BMC were added to the plates at a concentration of 5 x 105 cells/ml and incubated overnight with pools of 20-mer peptides (overlapping by 10 aa) spanning the entire sequence of HIV-1 clade B Gag (HXB2), Pol (HXB2), Env (MN), or Nef (BRU) (AIDS Research and Reference Reagent Program, Division of AIDS, National Institute of Allergy and Infectious Diseases (NIAID), National Institutes of Health) at 2 μg/ml. After washing, 1 μg/ml biotinylated anti-IFN- mAb 7-B6-1 (Mabtech) was added to the plates for 2 h at room temperature. Following another round of washing, streptavidin-conjugated alkaline phosphatase (Southern Biotechnologies) was added, and the plates were incubated at room temperature for an additional hour. Last, NBT/5-bromo-4-chloro-3-indolyl phosphate; Sigma-Aldrich) was added for color development. Individual cytokine-producing cells were counted manually and/or by the C.T.L. ImmunoSpot analyzer (Cellular Technologies). In addition, immune responses to HLA-restricted peptides (8–10 mers) from HIV-1, CMV, EBV, and influenza were also measured using the ELISPOT at a concentration of 10–5 M (13). PHA (Sigma-Aldrich) (5 μg/ml), was used as a positive control and unstimulated cells, i.e., medium alone, as the negative control. Responses were considered positive if there were 100 spot-forming cells (SFC)4/106 cells and twice as many SFC as in the nonstimulated control wells. PBMC were added to the ELISPOT plates at a concentration of 1 x 106 cells/ml and incubated with the same peptide pools as the BMC. The SFC was normalized to SFC per CD8+ T cells by using the percentage of CD8+ cells present in either breast milk or PBMC using flow cytometry.
Quantitation of HIV-1 RNA in plasma and breast milk
HIV RNA levels in the plasma and breast milk samples were determined by the Amplicor UltraSensitive HIV-1 Monitor version 1.5 assay (Roche Diagnostics Systems), according to the manufacturer’s protocol. We have previously demonstrated that this assay can reproducibly and reliably detect HIV in breast milk (14). In addition, we and others (15, 16, 17) have shown that breast milk RNA levels are at least 10- to 100-fold less than those found in the blood. The breast milk RNA levels in the HIV-infected women in this study were all <50 copies/ml.
Surface marker staining
Phenotypic characterization of BMC and PBMC used cell-surface markers CD3, CD4, CD8, CD57, CD62L, CD103, HLA-DR, CD45RO, CD45RA, CD27, CD28, CD38, CCR7, and CCR9 (BD Biosciences). Stained cells were acquired using a BD Calibur flow cytometer (BD Biosciences) and analyzed with FlowJo version 4.3.1 software (Tree Star). Our design was to run 50,000 gated T cells (based on CD3 staining) for each stained specimen. When CD3 was not used as a marker, we gated on CD8-staining cells.
MHC class I tetramer analysis
Enumeration of epitope-specific cells by MHC class I tetramer complexes was performed using either HLA B7 CMV pp65 TPRVTGGGAM, HLA A2 HIV Gag p17 SLYNTVATL, HLA A2 HIV Pol ILKEPVHGV, HLA A3 HIV Gag p17 RLRPGGKKK, or HLA B35 HIV Pol NPDDIVIYQY (kindly provided by L. Tussey, Merck, West Point, PA) tetramers. All HIV-specific MHC class I tetramers were synthesized by the NIAID Tetramer Facility (AIDS Research and Reference Reagent Program, Division of AIDS, NIAID, National Institutes of Health) and conjugated with PE. B7-CMV was kindly provided by S. Boppana (University of Alabama at Birmingham) and was conjugated to allophycocyanin. Tetramer titration and staining conditions were maximized by using epitope-specific PBMC from appropriate HLA seropositive and seronegative volunteers. For each analysis, a minimum of 10,000 events are represented.
Statistics
Comparison of the frequency and phenotype of T cells between breast milk and blood compartments were made by the nonparametric Mann-Whitney U test using Analyze-It software.
Results
Study population
BM cells and PBMC from 18 lactating women were analyzed for phenotypic differences, seven were HIV positive and 11 were HIV negative. Breast milk was collected between 2 days and 4.5 mo postpartum with most samples being collected within 1 wk of delivery. Median plasma viral loads for the HIV-positive women were <50 copies/ml, and the median CD4 count was 387 cells/mm3. All HIV-positive women were on antiretroviral therapy.
Comparison of the frequency of Ag-specific CD8+ T cells in breast milk and blood
Ag-specific IFN--secreting CD8+ T cell responses measured in breast milk and blood were normalized to CD8 SFC/105 T cells so that direct comparisons could be made. Most responses found in breast milk were significantly increased when compared with PBMC. These findings applied to CD8+ T cells stimulated with HIV-specific peptide pools (Fig. 1, A–C) as well as to those stimulated with HLA class I-restricted peptides from HIV, EBV, CMV, and influenza and matched for the particular individuals HLA class I (Fig. 1, D–F).
FIGURE 1. HIV- and non-HIV-specific immune responses as measured by the IFN- ELISPOT assay normalized to the number of CD8+ T cells. A–C, PBMC and BMC were stimulated with overlapping 20-mer peptide pools from Gag, Pol, Env, and Nef. D–F, Immune responses to HLA-restricted peptides (8–10 mers) from HIV, CMV, EBV, and influenza. A2-Flu (GILGFVFTL); A3-Flu (ILRGSVAHK); A3-EBV EBNA3A (RLRAEAQVK); A3-EBV BRLF1 (RVRAYTYSK); A11-EBV EBNA3B (IVTDFSVIK); A11-EBV BRLF1 (ATIGTAMYK); B7-CMV (TPRVTGGGAM), B8-EBV EBNA3A (FLRGRAYGL). Representative BMC and PBMC are taken from HIV-seropositive women. Note that the responses are plotted on a log scale since some of the differences were large. Responses were measured for both breast milk and blood for all Ags shown; thus, lack of a response signifies that response was not detected.
Next, we performed HLA class I tetramer staining in three women who had detectable responses to HIV and/or CMV. All displayed increased frequency of tetramer binding T cells in breast milk when compared with PBMC (Table I). This is consistent with our prior studies wherein we found that the tetramer frequency in blood was only 0.22% compared with 0.65% in breast milk (12).
Table I. Class I tetramer staining comparison between BMC and PBMC
The data for the B7-CMV tetramer was derived from the same volunteer used to detect B7-CMV-specific peptide responses in the ELISPOT assay (Fig. 1F). Of note, although increased responses are seen for breast milk cells in this volunteer with both IFN- ELISPOT assay (Fig. 1F, 2/3 non-HIV Ags) and tetramer (Table I), the differences are not as pronounced as those seen for the other volunteers.
Intracellular IFN- staining after PBMC stimulation obtained from another HIV-infected woman with the A2-SL9 peptide also demonstrated a higher frequency of IFN--secreting CD8+ T cells in breast milk (1.72%) when compared with PBMC (0.13%) (data not shown). Thus, with few exceptions, the vast majority of responses were detected at higher frequencies in breast milk.
Comparison of surface marker expression between BMC and PBMC
Given the differences in frequency of CD8+ Ag-specific T cells noted between BMC and PBMC, we were interested in determining whether the phenotype of these two populations of lymphocytes differed, especially since they represented lymphocytes from lymphoid and nonlymphoid compartments. When looking at standard lymphocyte surface markers, we noted that the median CD3+/CD8+ BMC percentages in both HIV positive and HIV negative were slightly greater when compared with PBMC (Table II). However, the total number of lymphocytes per milliliter present in breast milk is lower than PBMC (data not shown). The only significant difference when comparing HIV-positive to HIV-negative women was the finding of consistently lower percentage of CD4 T cells in the former group in PBMC (29.4% vs 54%, p = 0.012), and lower CD103 in BMC of HIV-positive women when compared with HIV negative (14.2% vs 32.2%, p = 0.003) (Table II). All other surface cell phenotypic markers were not statistically different when comparing HIV-positive to HIV-negative women.
Table II. Surface marker expression comparison between BMC and PBMC
Expression of CD103, an integrin directed to E-cadherin was at least an order of magnitude higher in BMC compared with PBMC in both HIV-positive and HIV-negative women (Table II and Fig. 2). Increased CD103 is suggestive of BMC CD8+ cell migration from epithelial compartments of the gastrointestinal tract. CCR9, another mucosal homing receptor, was also significantly higher in BMC of HIV-seronegative women when compared with their PBMC (Table II and Fig. 2).
FIGURE 2. Surface marker expression on CD8+ T cells from 18 lactating women. A, HIV-seropositive women (n = 7); B, HIV-seronegative women (n = 11). Surface marker expression on BMC vs PBMC is shown. Cells were gated by using the following parameters: forward scatter vs side scatter and CD3+/CD8+ or CD8+ T cells if CD3 was not performed. The median percentage of CD8+ T cells that stained with the given surface marker is plotted.
Reduced levels of CCR7 expression were detected on BMC (0.8% in HIV positive and 2.1% in HIV negative), whereas PBMC contained levels of expression as seen in other studies (28.2% in HIV positive and 52.2% in HIV negative) (10, 18). Furthermore, the large majority of T cells in breast milk from either HIV-seropositve or -seronegative women had a memory phenotype, as they expressed high levels of CD45RO, and reduced expression of CD45RA and the lymph node homing marker CD62L. The activation marker, HLA-DR, was highly expressed on the surface of breast milk-derived CD8+ T cells relative to PBMC in both HIV-positive and HIV-negative women (Fig. 2 and Table II). In contrast, breast milk from both HIV-positive and HIV-negative women and PBMC from HIV-positive women expressed high levels of another activation marker CD38. Levels of CD38 in PBMC of HIV-negative women were lower but not significantly different (Table II). Similar levels of CD57 expression, a marker suggestive of late memory T cells with little proliferative capacity (19, 20, 21) were observed when comparing the level of expression between BMC and PBMC from either HIV-seropositive or HIV-seronegative women (Table II and Fig. 2).
In short, the measured surface phenotypes on most T cell populations in the BMC samples from HIV-seronegative women differed significantly from the populations in the corresponding peripheral blood, with respect to all tested T cell markers except for CD38, CD57, CD4, and CD8 (Table II). The same trends were noted for BMC derived from HIV-positive women, but statistically significant differences were not as consistently observed most likely due to a small sample size.
Comparison of surface marker expression on Ag-specific CD8+ T cells from BMC and PBMC
The above data showed marked differences in the surface marker expression on whole populations of BMC vs PBMC. We next evaluated whether these differences were present in Ag-specific populations. Cells from two HIV-infected lactating women with known tetramer-positive responses were studied. Responses to B35Pol-NY9 were from volunteer 1 and responses to A2Pol-IV9 and B7-CMV from volunteer 2 (Table I). Similar to the finding with the whole T cell populations, differences were observed in the expression of CD45RA (lower in BMC from both HIV- and CMV-specific populations). HLA-DR also showed greater expression on the surface of BMC than PBMC. These differences occurred whether the Ag-specific cells were HIV or CMV specific (Fig. 3). We also determined the relative differentiation status of these Ag-specific cells by using the expression of the co-stimulatory receptors CD28 and CD27. These receptors are involved in the regulation of T cell activation and in the generation of Ag-primed cells, respectively, and have been used by others to distinguish between subsets of differentiated CD8+ T cells (4, 6). CD28 seems to be expressed at higher levels on the surface of BMC, whereas CD27 appears to be present at similar levels on both BMC and PBMC (Fig. 3).
FIGURE 3. Surface marker expression on Ag-specific cells from two HIV-infected women. Cells were gated on CD8+ tetramer + cells. HIV-1 Pol (aa 330–338, NPDIVIYQY) HLA B35 restricted staining, HIV-1 Pol (aa 309–317, ILKEPVHGV) HLA A2 restricted staining and CMV pp65 (aa 417–426, TPRVTGGGAM) HLA B7
Discussion
Memory T cells have been divided into multiple functional phenotypes on the basis of surface molecule expression, anatomic location, and immune function. Currently, there is no clear consensus on the pathways of memory/effector T cell differentiation, and several models have been proposed (2, 3, 6, 7, 22, 23, 24, 25, 26, 27). Studies in mice suggest that nonlymphoid tissues are preferentially populated by a subset of CD8+ T cells (9, 28). These cells express low levels of CCR7 and CD62L and are capable of producing effector cytokines immediately upon Ag stimulation; they have been referred to as "effector" memory T cells (Tem). In the present study, we characterized the phenotype of CD8+ T cells derived from breast milk from both HIV-infected and uninfected women. Overall, the phenotype of breast milk cells in these populations were remarkably similar, no doubt reflecting the fact all the HIV-infected women had normal levels of CD4 cells and were on highly active antiretroviral therapy. Using both ELISPOT and tetramer assays, we found that the frequency of Ag-specific CD8+ T cells in breast milk is significantly higher than PBMC. This is consistent with our previously published work as well as that of others (11, 12). Higher levels of Ag could not explain the higher frequency of Ag-specific cells, since in the case of HIV infection, breast milk contains much less HIV than plasma (12, 17, 29). In addition, compared with their counterparts in blood, breast milk CD8+ T cells lack expression of CCR7, CD62L, and CD45RA and express high levels of CD45RO. Breast milk T cells also express higher levels of the activation marker HLA-DR. Thus, the lactating breast is enriched for Ag-specific T cells distinct from those circulating in the blood, and these cells bear phenotypic hallmarks of effector memory cells that have been recently activated.
Expression of the co-stimulatory molecules CD27 and CD28 have also been associated with different stages of T cell differentiation (4, 6, 30). Sequential down-regulation of CD28 and then CD27 expression has been described during CD8+ T cell differentiation (4, 31). Our data on the expression of these markers for CMV-specific responses in PBMC of a lactating HIV-positive woman are consistent with that described by others (4, 27). CMV-specific cells express levels of CD27 and CD28 similar to those reported (4, 6), with concomitant expression of CD45RA showing as others have that these cells are in an intermediate to late stage of differentiation (4, 27). The low expression of CD45RA and high expression of HLA-DR on all Ag-specific breast milk cells (Fig. 3) is consistent with our data on whole BMC populations (Fig. 2 and Table II). Although we did not measure CCR7 on Ag-specific cells, it is unlikely that these cells would express this marker since very few of the total breast milk-derived population of CD8+
T cells expressed CCR7 (0.8% in HIV positive and 2.1% in HIV negative) (Table II and Fig. 2). Moreover, studies of Ag-specific T cells have demonstrated that greater than 70% of the Ag-specific CD8+ T cells do not express CCR7 (4, 7).
Recent studies have defined two subsets of effector memory (CD45RA–CCR7–) T cell subsets in humans, one expressing CD27 (CD27+) and one that does not (CD27–) (32). In our Ag-specific breast milk-derived CD8+ T cell populations, we have both of these populations represented (Fig. 3). In humans, the CD45RA–CD27+ subset is capable of cytolytic activity after in vitro stimulation (31). We have previously demonstrated that CD8+ breast milk-derived T cells were also capable of Ag-specific cytolysis after in vitro stimulation (12).
The expression of CD28 on breast milk Ag-specific cells was relatively high. According to some models of phenotypic evolution, expression of this marker is a sign of "earlier" differentiation and suggests that breast milk cells may be at an earlier stage in postthymic development than those cells found in the blood. Presumably this is due to less Ag in breast milk and/or exposure to other factors in the unique mucosal environment of the lactating breast. In addition, this may have something to do with the recent acquisition/homing of these cells into this compartment, given that breast milk is not produced before pregnancy. Given the limited number of Ag-specific samples analyzed, it is difficult to make definitive conclusions, and analysis of more samples is needed.
The origin of breast milk T cells is uncertain. Animal studies clearly demonstrate that B cells from Peyer’s patches migrate specifically to the lactating mammary gland (33, 34, 35, 36) and that pregnancy increases the mucosal vascular addressin MadCAM-1, which interacts with the gut homing receptor 47 (37). In humans, oral immunization with nonpathogenic Escherichia coli, produces type-specific IgA in breast milk but not in serum or saliva of lactating women, suggesting trafficking between breast and blood (38). There are few data on T cell migration between the intestine and the lactating breast. Labeling studies in rats and pigs demonstrate that T cells in the breast originate from the gut as well as blood (39, 40). We studied the expression of two mucosal markers CD103 (E7) and CCR9 in breast milk lymphocytes. CD103 is expressed on intraepithelial lymphocytes in the gut, genitourinary tract, and skin, as well as its increased expression in activated CD8+ cells (41). In contrast to PBMC where CD103 is expressed on very few lymphocytes (<2%), we found that it was expressed on a significant number of breast milk lymphocytes (32.2% in HIV negative and 14.2% in HIV positive), suggesting selective recruitment or retention of these cells. CCR9 is a chemokine receptor expressed on thymocytes, intestinal lymphocytes, and on discrete subsets of T cells that traffic to the intestine (42, 43). Expression of CCR9 on breast milk cells was significantly greater than that found in blood in HIV-negative women. Thus, expression of these mucosal homing receptors by CD8+ T cells from breast milk suggests that many of these cells may have originated from the gastrointestinal tract.
Taken together, these data demonstrate that breast milk is populated with CD8+ T cells that have an effector memory phenotype, consistent with studies in animals showing that a substantial portion of the T cell response to infection is focused on nonlymphoid tissue (1, 9, 44, 45, 46, 47). Study of breast milk cells provides a novel convenient method for the study of these cells in humans. Given the importance of milk in protecting progeny from infection, the enrichment of Ag-specific cells with an effector phenotype is teleologically appealing. The immunologic benefits of breast-feeding are well established (48), but the role of these effector memory T cells in the prevention of infection remains to be elucidated.
Disclosures
The authors have no financial conflict of interest.
Acknowledgments
We thank the volunteers for their participation, Barbara Corley and Shawn Dillon for breast milk procurement, Marion Spell and Tracy McGuire for help with the flow cytometric acquisition, Anju Bansal for insightful discussions, Linda Tussey for providing the B35 tetramer, Suresh Boppana for providing the B7 CMV tetramer and the Children’s Hospital Family Clinic.
Footnotes
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
1 This work was supported by grants from the National Institutes of Health (R01 HD-40777, HD-396110, AI-049126) and the Elizabeth Glaser Pediatric AIDS Foundation. G.M.A. is an Elizabeth Glaser Pediatric AIDS Foundation Scientist.
2 S.S. and M.K.G. contributed equally to this work.
3 Address correspondence and reprint requests to Dr. Grace M. Aldrovandi, Infectious Diseases #51, Children’s Hospital Los Angeles, 4650 Sunset Boulevard, Los Angeles, CA 90027. E-mail address: galdrovandi{at}chla.usc.edu
4 Abbreviation used in this paper: SFC, spot-forming cells.
Received for publication July 19, 2004. Accepted for publication December 20, 2004.
References
Masopust, D., V. Vezys, E. J. Usherwood, L. S. Cauley, S. Olson, A. L. Marzo, R. L. Ward, D. L. Woodland, L. Lefrancois. 2004. Activated primary and memory CD8 T cells migrate to nonlymphoid tissues regardless of site of activation or tissue of origin. J. Immunol. 172:4875.
Wherry, E. J., V. Teichgraber, T. C. Becker, D. Masopust, S. M. Kaech, R. Antia, U. H. von Andrian, R. Ahmed. 2003. Lineage relationship and protective immunity of memory CD8 T cell subsets. Nat. Immunol. 4:225.
Sallusto, F., D. Lenig, R. Forster, M. Lipp, A. Lanzavecchia. 1999. Two subsets of memory T lymphocytes with distinct homing potentials and effector functions. Nature 401:708.
Appay, V., P. R. Dunbar, M. Callan, P. Klenerman, G. M. Gillespie, L. Papagno, G. S. Ogg, A. King, F. Lechner, C. A. Spina, et al 2002. Memory CD8+ T cells vary in differentiation phenotype in different persistent virus infections. Nat. Med. 8:379.
Tomiyama, H., T. Matsuda, M. Takiguchi. 2002. Differentiation of human CD8+ T cells from a memory to memory/effector phenotype. J. Immunol. 168:5538.
Tussey, L. G., U. S. Nair, M. Bachinsky, B. H. Edwards, J. Bakari, K. Grimm, J. Joyce, R. Vessey, R. Steigbigel, M. N. Robertson, et al 2003. Antigen burden is major determinant of human immunodeficiency virus-specific CD8+ T cell maturation state: potential implications for therapeutic immunization. J. Infect. Dis. 187:364.
Champagne, P., G. S. Ogg, A. S. King, C. Knabenhans, K. Ellefsen, M. Nobile, V. Appay, G. P. Rizzardi, S. Fleury, M. Lipp, et al 2001. Skewed maturation of memory HIV-specific CD8 T lymphocytes. Nature 410:106.
Sabbaj, S., B. H. Edwards, M. K. Ghosh, K. Semrau, S. Cheelo, D. M. Thea, L. Kuhn, G. D. Ritter, M. J. Mulligan, P. A. Goepfert, G. M. Aldrovandi. 2002. Human immunodeficiency virus-specific CD8+ T cells in human breast milk. J. Virol. 76:7365.
Sabbaj, S., A. Bansal, G. D. Ritter, C. Perkins, B. H. Edwards, E. Gough, J. Tang, J. J. Szinger, B. Korber, C. M. Wilson, et al 2003. Cross-reactive CD8+ T cell epitopes identified in US adolescent minorities. J. Acquir. Immune Defic. Syndr. 33:426
Ghosh, M. K., L. Kuhn, J. West, K. Semrau, D. Decker, D. M. Thea, G. M. Aldrovandi. 2003. Quantitation of human immunodeficiency virus type 1 in breast milk. J. Clin. Microbiol. 41:2465.
Nduati, R., G. John. 1995. Breast milk transmission of HIV-1. NARESA Mongr Dec:1.
Semba, R. D., N. Kumwenda, D. R. Hoover, T. E. Taha, T. C. Quinn, L. Mtimavalye, R. J. Biggar, R. Broadhead, P. G. Miotti, L. J. Sokoll, et al 1999. Human immunodeficiency virus load in breast milk, mastitis, and mother-to-child transmission of human immunodeficiency virus type 1. J. Infect. Dis. 180:93.
Willumsen, J. F., M. L. Newell, S. M. Filteau, A. Coutsoudis, S. Dwarika, D. York, A. M. Tomkins, H. M. Coovadia. 2001. Variation in breastmilk HIV-1 viral load in left and right breasts during the first 3 months of lactation. AIDS 15:1896.
Chen, G., P. Shankar, C. Lange, H. Valdez, P. R. Skolnik, L. Wu, N. Manjunath, J. Lieberman. 2001. CD8 T cells specific for human immunodeficiency virus, Epstein-Barr virus, and cytomegalovirus lack molecules for homing to lymphoid sites of infection. Blood 98:156.
Brenchley, J. M., N. J. Karandikar, M. R. Betts, D. R. Ambrozak, B. J. Hill, L. E. Crotty, J. P. Casazza, J. Kuruppu, S. A. Migueles, M. Connors, et al 2003. Expression of CD57 defines replicative senescence and antigen-induced apoptotic death of CD8+ T cells. Blood 101:2711.
d’Angeac, A. D., S. Monier, D. Pilling, A. Travaglio-Encinoza, T. Reme, M. Salmon. 1994. CD57+ T lymphocytes are derived from CD57– precursors by differentiation occurring in late immune responses. Eur. J. Immunol. 24:1503
Kern, F., E. Khatamzas, I. Surel, C. Frommel, P. Reinke, S. L. Waldrop, L. J. Picker, H. D. Volk. 1999. Distribution of human CMV-specific memory T cells among the CD8pos. subsets defined by CD57, CD27, and CD45 isoforms. Eur. J. Immunol. 29:2908
Ahmed, R., D. Gray. 1996. Immunological memory and protective immunity: understanding their relation. Science 272:54.
Kaech, S. M., E. J. Wherry, R. Ahmed. 2002. Effector and memory T-cell differentiation: implications for vaccine development. Nat. Rev. Immunol. 2:251.
Kaech, S. M., J. T. Tan, E. J. Wherry, B. T. Konieczny, C. D. Surh, R. Ahmed. 2003. Selective expression of the interleukin 7 receptor identifies effector CD8 T cells that give rise to long-lived memory cells. Nat. Immunol. 4:1191.
Seder, R. A., R. Ahmed. 2003. Similarities and differences in CD4+ and CD8+ effector and memory T cell generation. Nat. Immunol. 4:835.
Lanzavecchia, A., F. Sallusto. 2002. Progressive differentiation and selection of the fittest in the immune response. Nat. Rev. Immunol. 2:982.
Appay, V., S. L. Rowland-Jones. 2004. Lessons from the study of T-cell differentiation in persistent human virus infection. Semin. Immunol. 16:205
Reinhardt, R. L., A. Khoruts, R. Merica, T. Zell, M. K. Jenkins. 2001. Visualizing the generation of memory CD4 T cells in the whole body. Nature 410:101.
Rousseau, C. M., R. W. Nduati, B. A. Richardson, M. S. Steele, G. C. John-Stewart, D. A. Mbori-Ngacha, J. K. Kreiss, J. Overbaugh. 2003. Longitudinal analysis of human immunodeficiency virus type 1 RNA in breast milk and of its relationship to infant infection and maternal disease. J. Infect. Dis. 187:741.
Hamann, D., M. T. Roos, R. A. van Lier. 1999. Faces and phases of human CD8 T-cell development. Immunol. Today 20:177.
Hamann, D., P. A. Baars, M. H. Rep, B. Hooibrink, S. R. Kerkhof-Garde, M. R. Klein, R. A. van Lier. 1997. Phenotypic and functional separation of memory and effector human CD8+ T cells. J. Exp. Med. 186:1407.
Rufer, N., A. Zippelius, P. Batard, M. J. Pittet, I. Kurth, P. Corthesy, J. C. Cerottini, S. Leyvraz, E. Roosnek, M. Nabholz, P. Romero. 2003. Ex vivo characterization of human CD8+ T subsets with distinct replicative history and partial effector functions. Blood 102:1779.
Roux, M. E., M. McWilliams, J. M. Phillips-Quagliata, M. E. Lamm. 1981. Differentiation pathway of Peyer’s patch precursors of IgA plasma cells in the secretory immune system. Cell. Immunol. 61:141
Tanneau, G. M., L. Hibrand-Saint Oyant, C. C. Chevaleyre, H. P. Salmon. 1999. Differential recruitment of T- and IgA B-lymphocytes in the developing mammary gland in relation to homing receptors and vascular addressins. J. Histochem. Cytochem. 47:1581
Goldblum, R. M., S. Ahlstedt, B. Carlsson, L. A. Hanson, U. Jodal, G. Lidin-Janson, A. Sohl-Akerlund. 1975. Antibody-forming cells in human colostrum after oral immunisation. Nature 257:797.
Manning, L. S., M. J. Parmely. 1980. Cellular determinants of mammary cell-mediated immunity in the rat. I. The migration of radioisotopically labeled T lymphocytes. J. Immunol. 125:2508.
Salmon, H.. 2000. Mammary gland immunology and neonate protection in pigs. Homing of lymphocytes into the MG. Adv. Exp. Med. Biol. 480:279.
Agace, W. W., J. M. Higgins, B. Sadasivan, M. B. Brenner, C. M. Parker. 2000. T-lymphocyte-epithelial-cell interactions: integrin (E)(CD103)(7), LEEP-CAM and chemokines. Curr. Opin. Cell Biol. 12:563.
Zabel, B. A., W. W. Agace, J. J. Campbell, H. M. Heath, D. Parent, A. I. Roberts, E. C. Ebert, N. Kassam, S. Qin, M. Zovko, et al 1999. Human G protein-coupled receptor GPR-9-6/CC chemokine receptor 9 is selectively expressed on intestinal homing T lymphocytes, mucosal lymphocytes, and thymocytes and is required for thymus-expressed chemokine-mediated chemotaxis. J. Exp. Med. 190:1241.
Svensson, M., J. Marsal, A. Ericsson, L. Carramolino, T. Broden, G. Marquez, W. W. Agace. 2002. CCL25 mediates the localization of recently activated CD8+ lymphocytes to the small-intestinal mucosa. J. Clin. Invest. 110:1113.
Klonowski, K. D., K. J. Williams, A. L. Marzo, D. A. Blair, E. G. Lingenheld, L. Lefrancois. 2004. Dynamics of blood-borne CD8 memory T cell migration in vivo. Immunity 20:551
La Gruta, N. L., S. J. Turner, P. C. Doherty. 2004. Hierarchies in cytokine expression profiles for acute and resolving influenza virus-specific CD8+ T cell responses: correlation of cytokine profile and TCR avidity. J. Immunol. 172:5553.
Sun, J. C., M. J. Bevan. 2004. Cutting edge: long-lived CD8 memory and protective immunity in the absence of CD40 expression on CD8 T cells. J. Immunol. 172:3385.
Wherry, E. J., J. N. Blattman, K. Murali-Krishna, R. van der Most, R. Ahmed. 2003. Viral persistence alters CD8 T-cell immunodominance and tissue distribution and results in distinct stages of functional impairment. J. Virol. 77:4911
Fituch, C., K. Palkowetz, A. Goldman, R. Schanler. 2004. Concentrations of IL-10 in preterm human milk and in milk from mothers of infants with necrotizing enterocolitis. Acta Paediatr. 93:1496(Steffanie Sabbaj, Mrinal )