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Intestinal T Cells Differentiate and Rearrange Antigen Receptor Genes In Situ in the Human Infant¹

Amanda M. Williams^2,*, Paul W. Bland^,, Anne C. Phillips, Susan Turner^*, Trevor Brooklyn^*, Gabriel Shaya^*, Richard D. Spicer and Christopher S. J. Probert^*

Departments of ^* Clinical Science at South Bristol and Clinical Veterinary Science, University of Bristol, and Department of Paediatric Surgery, Bristol Royal Hospital for Children, Bristol, United Kingdom; and Department of Clinical Immunology, University of Gothenburg, Gothenburg, Sweden

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion References

 
Intestinal Ag exposure during neonatal life influences appropriate adult immune responses. To define the mechanisms shaping the T cell repertoire during this period, we examined T cell differentiation and receptor diversity in the intestine of human infants. Developmental phenotypes of intraepithelial and lamina propria intestinal T cells from infants aged 1 day to 2 years were assessed ex vivo by flow cytometry and in situ by triple-fluorescent immunohistochemistry. Gene recombination-specific enzymes were assessed by PCR. TCR -chain V region gene diversity was determined by sequencing. Several different early lineage T cell populations were present neonatally: CD3⁺4^–8^– T cells were present at birth and numbers decreased during the neonatal period; CD3⁺4⁺8⁺ T cells were present in low numbers throughout infancy; and CD3⁺4⁺8^– or CD3⁺4^–8⁺ T cells increased with age. Very early lineage T cells, CD3^–2^–7⁺ and CD3^–2⁺7⁺, were present neonatally, but were essentially absent at 1 year. Most lamina propria T cells differentiated rapidly after birth, but maturation of intraepithelial T cells took place over 1 year. Intestinal samples from infants less than 6 mo old contained transcripts of T early and TdT, and 15 of 19 infant samples contained mRNA for RAG-1, some coexpressing RAG-2. TCR -chain repertoires were polyclonal in infants. Immature T cells, pre-T cells, and genes involved in T cell recombination were found in the intestine during infancy. T cell differentiation occurs within the neonatal human intestine, and the TCR repertoire of these developing immature T cells is likely to be influenced by luminal Ags. Thus, mucosal T cell responsiveness to environmental Ag is shaped in situ during early life.

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion References

 
The ontogeny of human intestinal T cells is poorly understood. It has been suggested that some undergo extrathymic development, and there is some evidence that the repertoire, particularly for T cells, is shaped in infancy. However, the complexity of the TCR repertoire in early life has received little attention.

In mice, extrathymic T cell maturation of a subpopulation of intestinal intraepithelial lymphocytes (IEL)³ has been proposed and much evidence to substantiate this theory has accumulated in recent years. The majority of intestinal intraepithelial T lymphocytes are CD8⁺ and express the TCR. In mice, but not in human adults, many of these CD8⁺ IEL express CD8 as an homodimer (1, 2), rather than as an heterodimer. Intestinal T cells of the TCR CD8 phenotype can express TCR -chains capable of reacting with self-superantigens (1, 2, 3), but any T cells that are likely to respond in a self-reactive manner would normally undergo clonal deletion in the thymus. As TCR CD8 cells are found exclusively in the intestinal epithelial compartment, it has been suggested, therefore, that they have developed in the intestine (3), thus evading negative selection and clonal deletion in the thymus. Further evidence for intestinal T cell development in mice is the presence of mRNA for recombination-activating proteins RAG-1 and RAG-2 (1, 4, 5, 6, 7). In contrast, a recent study indicates that recombination events in mouse intestinal mucosal T cells are confined to athymic individuals and occur in the draining mesenteric lymph nodes (8). However, even in mice, the extent to which mucosal T cells differentiate in situ during the neonatal stage is still unclear.

In humans, evidence for extrathymic maturation in the intestine is limited. Immature T cells (9, 10) and TCR CD8 cells (11) have been found in human fetal intestine, but their presence in the intestine after birth has not been reported. Expression of RAG mRNA has been shown in human adult IEL (12), but recombination-activating proteins are not exclusive to T cells (7, 13, 14). Conversely, another gene associated with TCR rearrangement, TdT, was absent from the intestine of human adults (15). T early (TEA) is expressed in immature T cells at the point when double-negative (DN) thymocytes become immature single-positive (SP) T cells (16), and is required for TCR recombination (17). TEA protein and concordant expression of TEA mRNA was reported in the intestine of human fetuses (9), but has not been investigated postnatally.

The TCR and TCR repertoire of IEL in human adults is oligoclonal (18, 19, 20) and it is assumed that this reflects the need to tightly regulate the immune response in the intestine. However, Holtmeier et al. (20) have reported a polyclonal repertoire in midgestation, retained after delivery, but becoming oligoclonal during childhood. Regulation of this restricted repertoire is not understood, but if IEL undergo TCR rearrangement in the intestine, then it is likely that intestinal Ags present during rearrangement are important in shaping the repertoire. Therefore, an understanding of where and when T cells mature will help us to understand the factors driving the development of the oligoclonal repertoire evident in adulthood.

To determine whether T cell maturation occurs in the intestine of human infants, we have investigated the phenotype of T cells and their precursors, the expression of genes and proteins associated with TCR rearrangement, and the repertoire of TCR T cells.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
Patients

Intestinal tissue was obtained from surgical resection specimens. Only normal tissue close to the resection margins was studied, so that any disease process leading to the surgery did not influence the results (Table I). All experiments were conducted blind and patient age and sample information was retrieved from the study database at the end of the study. Fresh tissue was obtained for the work with informed parental consent. All aspects of this research were approved by the Research Ethics Committee for United Bristol Healthcare Trust.

Fresh tissue was collected on ice in RPMI 1640 (Sigma-Aldrich, Poole, Dorset, U.K.). The mucosa was dissected free of underlying tissue and two 5 mm x 10 mm pieces were mounted in Tissue-Tek OCT compound (RA Lamb, Sussex, U.K.) and frozen in liquid nitrogen-cooled isopentane for immunohistochemistry. Three match-head-size pieces of mucosa were stored in 1 ml of TRIzol (Invitrogen Life Technologies, Paisley, U.K.), for mRNA extraction within 1 mo. The remaining mucosa (1–2 cm²) was used for the isolation of lymphocytes, as previously described (21).

Isolated IEL and lamina propria lymphocytes (LPL) were passed through 70-µm sieves and purified over discontinuous Percoll gradients (Amersham, Bucks, U.K.) (40:60%). The cell count and viability was determined by trypan blue exclusion. Aliquots of each fraction were analyzed by flow cytometry and 10⁶ cells were resuspended in 1 ml of TRIzol for mRNA isolation.

Flow cytometry

All Abs were titrated using PBLs for use with a maximum of 10⁷ cells in a staining volume of 50 µl. Abs were supplied by Caltag-Medsystems (Towcester, U.K.), unless stated otherwise. Cell aliquots (50–100,000) were triple-stained with combinations of Abs against CD4 (FITC, IgG1; BD Biosciences, Oxford, U.K.), CD8 (R-PE, IgG2a), CD3 (tricolor, IgG2a), CD2 (FITC, IgG1), CD5 (R-PE, IgG1), and CD7 (R-PE, IgG2a). Isotype controls were FITC-mouse IgG1 and R-PE mouse IgG2a (DAKO, Cambridgeshire, U.K.) and analyzed using a FACSCalibur Flow Cytometer (BD Biosciences). Twenty thousand CD3⁺ events/sample were analyzed using CellQuest Pro Software (BD Biosciences).

RNA isolation and cDNA synthesis

RNA was extracted from whole fresh mucosa and from fractionated IEL and LPL using TRIzol according to the manufacturer’s protocol. Reverse transcription was performed using Expand Reverse Transcriptase (Roche, Sussex, U.K.) according to the manufacturer’s protocol. Integrity of the cDNA was confirmed by PCR amplification of a housekeeper gene, GAPDH, and visualized on a 2% agarose gel stained with ethidium bromide. The cDNA was kept at –20°C for later use.

Detection of transcripts from genes encoding TdT, TEA, RAG-1, and RAG-2

Stored cDNA was used as a template for four sets of PCR amplifications designed to identify transcripts of genes that are associated with germline recombination and rearrangement of the TCR gene locus. Primer pairs were used to investigate the expression of mRNA for TdT (15): TCAGAGTTCTGAAACCCATCCT (antisense) and AGACTCCACCAATTGCTGTACA (sense); TEA: GGCAGACAGACTTGTCACTGGAT (antisense) and GGGACACTCCATGGTGTTGTTGTTG (sense); RAG-1 (12): CTTGGCTTTCCAGAGAGTCC (antisense) and TGGATCTTTACCTGAAGATG (sense); and RAG-2 (12): CATCATCTTCATTATAGGTGTC (antisense) and TGGAAGCAACATGGGAAATG (sense). The same PCR conditions were used for all sets of PCR. A total of 1–3 µl of cDNA was amplified using appropriate primer pairs (10 pmol of each), 0.2 mM dNTPs and 0.25 U of TaqDNA polymerase (Roche). Each reaction consisted of 32 cycles: 94°C for 20 s, 55°C for 30 s, and 72°C for 60 s. The last cycle was followed by a final extension period of 7 min at 72°C and then chilled to 4°C. After each set of reactions, the PCR products were visualized on a 2% agarose gel as described previously.

TdT, TEA, and RAG-1 products were sequenced. PCR products were cloned directly into TA-cloning vector (Invitrogen Life Technologies) and competent Escherichia coli transformed and grown overnight. White colonies were picked, subcultured, and plasmid DNA extracted using the QIAprep Spin Miniprep kit (Qiagen, Crawley, U.K.). The plasmids were screened for appropriately sized inserts using EcoRI restriction enzyme (Roche). Subsequently, DNA sequencing was performed using a T7 primer, by The Sequencing Service (School of Life Sciences, University of Dundee, Dundee, Scotland; www.dnaseq.co.uk). The derived nucleotide sequence was compared with the published sequences for TdT, TEA, and RAG-1 using the nucleotide basic local alignment search tool (BLAST) search facility provided by the National Center for Biotechnology Information (NCBI) (www.ncbi.nlm.nih.gov, National Center for Biotechnology Information, Public Database of Molecular Biology Information. NCBI, U.S. National Library of Medicine, 8600 Rockville Pike, Bethesda, MD 20894).

Assessment of TCR repertoire

TCR -chain V region (TCRBV) 4, 6, and 12 genes were sequenced in 11 samples from eight children. These are reported to be the three most commonly used TCR -chains (22). Stored cDNA was used as a template for PCR amplifications, and each specific TCRBV PCR was conducted on all patient samples simultaneously. No contaminating sequences were found between unrelated patient samples. A primer specific to a conserved region of the TCR -chain C region (TCACCCACCAGCTCAGCT) was used with a second primer specific for the appropriate TCRBV: TCRBV4 (CCACATATGAGAGTGGATTTGTCA), TCRBV6 (CAGTGATCGGTTCTCTGCAGA, primed for; 6s3, 6s4, 6s5, 6s7, 6s11, 6s14), TCRBV12 (TACTGACAAAGGAGAAGTCTCAGA, primed for; 12s2, 12s4).

For each reaction, a 1–3 µl template was amplified using appropriate primer pairs (2 nM), 1 mM dNTPs (Roche), and 0.4 U TaqDNA polymerase (Promega, Southampton, U.K.). PCR amplification of all TCRBVs was performed at 94°C for 20 s, 55°C for 30 s, and 72°C for 60 s, with a final extension period of 7 min at 72°C and then chilled to 4°C. For amplification of TCRBV6, the reaction consisted of 40 cycles; for TCRBV4 and TCRBV12 the reaction consisted of 36 cycles. After each set of reactions, the PCR products were resolved on 2% agarose gels. Appropriately sized bands were cut from the gel using sterile scalpel blades and purified using the QIAquick Gel Extraction kit (Qiagen) before cloning in the pCR2.1 TOPO TA cloning kit (Invitrogen Life Technologies), and sequencing as described previously. Derived sequences were compared with published sequences from the NCBI database, as previously described.

Immunohistochemistry and image analysis

Frozen sections were cut at 5 µm, air-dried for 1 h, and then fixed for 10 min in precooled acetone at 4°C. Rehydrated sections were blocked with 10% normal goat and 10% human serum in PBS. Primary Abs were applied at room temperature for 3 h, and directly conjugated secondary Abs for 1 h at room temperature, with PBS washes before and after the secondaries. For double and triple labeling, all primary Abs were applied simultaneously. Sections were mounted in Vectashield (Vector Laboratories, Peterborough, U.K.). Controls included substitution of primary Abs with isotype-matched Ag-irrelevant Abs and incubations with secondary Ab only.

For both large and small bowel immunohistochemical sections, cell counting was performed using 5–7 crypt or crypt-villus units from three to four distinct well-orientated representative fields of view, at x200 or x400 magnification on a Leica DMR microscope with a triple filter and mercury lamp, using the Image Pro-Plus capture and analysis software (Media Cybernetics U.K., Berkshire, U.K.). Cell counts were made from measured areas of whole intestine and numbers of IEL or LPL found within the measured area were equated to cells/mm².

Statistics

Correlations between different cell types and age were assessed by regression analysis using Pearson correlation (r) and are indicated where appropriate in the text. Statistical significance was defined as p < 0.05. Data derived from flow cytometry analysis was separated into groups on a preweaning/postweaning basis to allow us to conduct two sample unpaired t tests and thus give a reflection of changes due to weaning. We proposed that 9 mo (6 mo after the usual onset of weaning) would be an appropriate time point for us to assume good exposure to food Ag had occurred and some adaptation of the T cells would be evident. Data derived from flow cytometry analysis was subjected to two sample unpaired t tests. Significance was defined as p < 0.05 by Student’s t test.

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
Precursor T cells are present in the intestinal epithelium and lamina propria during the postnatal period and in declining numbers with age

Aliquots of isolated intestinal lymphocytes from five small bowel samples (ages 6 days-12 mo, median = 2.5 mo) and 11 colonic samples (ages 1 day-4.75 years, median = 9 mo) were analyzed by flow cytometry. The distribution of CD4 and CD8 within the CD3⁺ gate of IEL and LPL fractions of small and large intestine was determined (Table II).

Cells coexpressing CD4 and CD8 (double-positive (DP) cells) accounted for up to 10% of CD3⁺ cells in both IEL and LPL. The proportion of DP cells changed little with age.

The proportion of total SP cells (CD3⁺4⁺ plus CD3⁺8⁺ cells) increased with age. There was a significant negative correlation between the proportion of DN cells and total SP cells in both the small (r = –0.99, p < 0.0001, IEL; and r = –0.97, p < 0.006 LPL) and large intestine (r = –0.99, p < 0.0001, IEL; and r = –0.89, p < 0.0003 LPL). When the CD8⁺ fraction of SP cells was studied, there was a strong negative correlation between CD3⁺8⁺ cells and CD3⁺ DN cells in the IEL compartment. This was significant in the large intestine (age <5 years, n = 11, r = –0.65, p < 0.03) but not in the small intestine (ages <5 years, n = 5, r = –0.85, p = 0.067) although the study may be underpowered. This indicates that in the epithelium, DN cells decrease concordant with increased CD3⁺8⁺ cells, most probably due to recruitment of CD3⁺8⁺ cells to the IEL compartment.

The increase in CD4⁺ and CD8⁺ SP cells, and decrease in DN cells, with age, was confirmed by observations of a single patient (D) from whom two large bowel samples (PT7 and PT13) were obtained at 6 and 9 mo of age. Between these ages this patient displayed an increased proportion of CD3⁺4⁺ SP IEL (from 3.2 to 20.1%), an increased proportion of CD3⁺8⁺ SP IEL (from 1.8 to 49.0%) and LPL (from 14.0 to 32.3%), and a decreased proportion of CD3⁺ DN IEL (from 94.8 to 27.5%) and LPL (from 30.3 to 13.1%).

To further investigate the expression of CD4 and CD8, immunohistochemistry studies were performed. Cell counting was conducted on five ileal samples (ages 6 days-12 mo) and eight colonic samples (ages 1 day-18 mo) (Table III). These studies also showed that CD4⁺ cells were often as numerous as CD8⁺ cells, indeed CD4⁺ cells accounted for 20–77% of T cells in the ileal mucosa and 54–78% in the colonic mucosa.

Flow cytometry demonstrated a large proportion of DN cells and that most were lymphocytes. Those DN cells that were CD3⁺ have been described above. However, there appeared to be other cells of similar light scatter characteristics that did not stain for CD3, suggesting that these may be CD3^– lymphocyte precursors. To clarify this, expression of CD2, CD5, and CD7 were assessed in the IEL and LPL from seven subjects (age 1 day-27 mo, Table II). In both compartments of the small and large intestine, four quadrants were defined within the CD3^+/– gated lymphocytes based on their expression of CD2 and CD5, or CD2 and CD7.

CD3^–7⁺2⁺ and CD3^–7⁺2^– populations, which are both most likely early T cell precursors, were at highest levels in tissues from subjects aged 9 mo or less, dropping to almost undetectable levels in subjects aged >9 mo (Table II). CD3^–CD7⁺CD2^– cells (the phenotype of bone marrow-derived T cells or NK precursors) were found in IEL and LPL in small and large intestine. CD5⁺CD2⁺ cells were also present in both immunologic compartments in the small and large intestine. Most of these cells also stained positively for CD3, but up to 11% of the CD5⁺CD2⁺ lymphocytes were CD3^–, and these were likely to be immature T cells and unlikely to be precursors of either NK or B cells. For each individual sample, the proportion of CD5⁺CD2⁺ cells was greater in the LPL than the IEL fraction. The proportion of CD5⁺CD2⁺CD3^– lymphocytes tended to decrease with age, with the maximum expression observed in the 1-day-old child (O), accounting for 6.5% of CD3^– lymphocytes in the LPL fraction (1.6% of total lymphocytes), and 2.9% of CD3^– lymphocytes in the IEL fraction (2.2% of total lymphocytes). All intestinal regions showed an increase in proportion of the CD3^–5⁺2^– subpopulation, possibly B cells, from <9 mo to >9 mo. The presence of immature cells was also investigated by triple immunofluorescence histology (Table III, Fig. 1). CD5 is expressed by both T and B cells, CD7 by T, NK, and their precursors. Thus, only T cell precursors will express both CD5 and CD7, whereas the presence of CD3, CD16 and CD19 would indicate mature T, NK, and B cells, respectively. In the small intestine (n = 6), SP CD7⁺ cells were found in both the epithelium (38–165/mm²) and the lamina propria (241–725/mm²), while SP CD5⁺ cells were less numerous (42–249/mm²) in the lamina propria and essentially absent from the epithelium. In the colon (n = 9), SP CD7⁺ cells were found in both the epithelium (0–146/mm²) and the lamina propria (77–938/mm²), while SP CD5⁺ cells were less numerous (12–192/mm²) in the lamina propria and, again, essentially absent from the epithelium. CD5⁺7⁺ DP cells were present in every lamina propria sample (2–540/mm² in the ileum, 16–328/mm² in the colon), although very few CD5⁺7⁺ DP cells were found in the epithelium (0–20/mm² in the ileum, 0–57/mm² in the colon).

View larger version (68K): [in this window] [in a new window]   FIGURE 1. CD3– lymphocytes in the human infant intestine express surface membrane Ags of immature T cells. Ileal tissues from patients aged 6 days (a) and 3.9 mo (b) underwent immunohistochemistry with FITC-conjugated CD3, CD16a, and CD19, Texas Red-conjugated CD5 (CD3–5+, red), and 7-amino-4-methylcoumarin-3-acetic acid (AMCA)-conjugated CD7 (CD3–7+, blue). Colonic tissues from patients aged 4 mo (c) and 9 mo (d) underwent immunohistochemistry with AMCA-conjugated CD3, CD16a and CD19, FITC-conjugated CD5 (CD3–5+, green), and Texas Red-conjugated CD7 (CD3–7+, red). Scale bars indicate 100 µm. CD3–7+ pre-T cells were observed in the lamina propria, and occasionally in the epithelium (arrowed). Occasionally pre-T cells were observed coexpressing CD5 and CD7 but without expression of CD3, CD16, or CD19. These cells are the phenotype of bone marrow-derived precursors. For the orientation of villi and crypts, Lu indicates the luminal surface.

The machinery required for genetic recombination exists in the intestinal mucosa of human infants, but wanes with age

PCR products of the appropriate size for TEA were obtained from 5 of 11 cDNA samples (age 3 days-18 years) (Table IV). All samples from infants aged 6 mo or less contained TEA (n = 5). The identity of the cDNA was confirmed by cloning and sequencing.

These data suggest that TEA and TdT genes are transcribed in the intestine during the first 18 mo of life, but are unlikely to be transcribed later. All samples from infants aged 6 mo or less contained transcripts for TEA and TdT. Seven samples were obtained from the small intestine, five contained the transcripts of interest. Too few large bowel samples were studied to conclude that there were fewer transcripts than in the small bowel.

Six samples used in the TdT and TEA studies were also analyzed for RAG transcript expression (Table IV). All contained transcripts for RAG-1 or RAG-1 and RAG-2. In total, 19 cDNA samples were analyzed and RAG-1 was found in 15, while RAG-2 was only found in 5. Concurrent expression of RAG-1 and RAG-2 was found in 4 samples.

Although TEA expression is thought to be specific for T cells, TdT and RAG may be expressed by non-T cells. In additional experiments, intestinal expression of TdT protein was investigated by immunohistochemistry in seven infants aged 18 mo or less (n = 7). Thymus tissue was a positive control and an isotype-matched Ab the negative control. TdT protein was detected in four of four small intestinal samples from three different infants (A-cyst, A-Ileum, B and L) and two of four large intestinal samples from four different infants (E and F positive, L and I negative). TdT protein was expressed mostly by cells in the lamina propria (Fig. 2, c and d), at least some of which were CD3⁺ (Fig. 2e), and by the occasional cell in the epithelium.

View larger version (147K): [in this window] [in a new window]   FIGURE 2. TdT protein is expressed in the human infant intestine. TdT protein expression was investigated by immunohistochemistry in small intestinal samples from patients aged 6 days to 4 mo and large intestinal samples from patients aged 2–18 mo. An isotype control Ab (a) was used as a negative control in all tissues. Thymus was used as positive control tissue (b). TdT protein was expressed by cells in the infant small intestine (four of four positive) and large intestine (two of four positive). Examples of TdT+ immunohistochemistry are shown for the small intestine (aged 6 days (c), and 3.9 mo (d), TdT+-DAB, brown) and large intestine (e, TdT-FITC, CD3-Texas Red). TdT protein expression was observed by cells in the lamina propria and occasionally in the epithelium. Although some cells displayed a clear nuclear distribution of TdT (filled arrow), many expressed TdT outside the nucleus. This is consistent with the translation of TdT protein and protein folding in the Golgi and endoplasmic reticulum before translocation to the nucleus where it is required for TCR recombination. Occasionally CD3+ cells were TdT+ (open arrow), but the majority of TdT+ cells were CD3–. Scale bars indicate 100 µm.

TCR -chain junctional sequences were obtained from three different families of TCRBV: TCRBV4, TCRBV6, and TCRBV12 (Table V). Initially, cDNA derived from small or large bowel mucosa whole tissue was used as the template. Subsequently, for two small intestinal samples and three large intestinal samples, cDNA from separated LPL and/or IEL was used. Two samples were obtained from a 6-day-old child (patient A) with an ileal duplication cyst. This allowed the effect of luminal contents on the small intestinal TCR repertoire to be determined as the cyst lay parallel to, but not in continuity with, the ileum. Small and large intestinal samples were obtained from a 2- and 1/4-mo-old infant (patient E) undergoing an ileostomy reversal. As this was created at birth, the colon had never been exposed to food Ags. Two colon samples were obtained from one child (patient D) separated by a 3-mo interval; samples were taken at 6 and 9 mo of age. Over 800 junctional regions were sequenced. The data presented are derived from samples where at least 20 junctional regions were sequenced, unless the first 10 sequences for a particular sample were identical.

In the large intestine, few clonal expansions of TCRBV6 and TCRBV12 were evident in the mucosal samples (Table V). Of the 21–27 sequences obtained per sample, most were different and six or fewer clones were found more than once. Occasionally a mucosal tissue sample displayed a more restricted TCRBV6 or TCRBV12 repertoire. However, when a corresponding LPL or IEL sample was studied, very few common clones were found and overall the repertoire of the patient was polyclonal. The TCR repertoire did not appear to be more restricted with increasing age, at least in the infants studied (aged 0–18 mo). In the oldest infant (18-mo-old, I) no TCRBV12 clonal expansions were found in LPL, and no TCRBV6 clonal expansions were found in IEL and LPL. Only one TCRBV6 sequence was found to be common, accounting for 4 of 24 clones from the mucosa and 1 of 21 clones from the IEL fraction. For comparison, TCRBV6 was studied in fractionated IEL from a child aged almost 5 years and at this age the TCRBV6 repertoire of IEL was more restricted, with 7 different sequences used by 24 clones (29% unique). Interestingly, the sample from the colon that had not been exposed to the fecal stream (E) was indistinguishable from the others; 17 of 21 sequences were different and just three clones appeared more than once (80% unique). Thus, although this child displayed a monoclonal TCRBV4 repertoire in its small intestine mucosa (E, Table V), the large intestinal mucosa displayed a polyclonal TCRBV4 repertoire, consistent with other large intestinal samples, despite being isolated from the normal flow of luminal Ags at birth.

Using TCRBV4, similar results were obtained: 19–29 sequences were obtained per sample and most samples were polyclonal with six or fewer clones found more than once. Some evidence of clonal restriction was found in the TCRBV4 repertoire of two samples. One mucosal sample (K, 12 mo, 38% unique) and one fractionated LPL sample (F, 9 mo, 27% unique) displayed a more restricted TCR repertoire. Of the two patients from which LPL fractions had been isolated, patient F had no sequences shared between the mucosa and the LPL fraction, and patient D had only one sequence common to the mucosal and LPL fractions which was used by only three clones. When the fractionated LPL samples were compared, the LPL from the older child (F, 9 mo) displayed a more restricted TCRBV4 repertoire, 27% of clones were unique, compared with the LPL from patient D (6 mo) in whom 80% were unique. When the sequences obtained from patient D at 6 and 9 mo were compared, only one sequence from the first LPL fraction, used by 1 of 26 clones, was found to be retained in the mucosa at 9 mo, and was slightly expanded accounting for 4 of 28 clones. In fact, the mucosal samples from patient D displayed a slightly more restricted TCRBV4 repertoire at 9 mo than at 6 mo, 50 and 60% unique, respectively.

The frequency with which each joining gene was used was determined from the 764 TCR sequences derived from the samples described above. The most commonly used TCRBJ segment was J2s3, followed by J2s7, J2s1, and J1s1 (Fig. 3). This is similar to that described for J usage in adult PBLs (22).

View larger version (39K): [in this window] [in a new window]   FIGURE 3. TCRBJ usage in human infant intestine is similar in T cells using TCRBV4, 6, and 12. The frequency with which each joining gene was used was determined from 764 TCR sequences. The most commonly used TCRBJ segment was J2s3, followed by J2s7, J2s1, and J1s1. This is similar to that reported for human adult PBLs.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
This study provides the first evidence that, in the human infant, differentiation of epithelial and mucosal T cells takes place in the intestine over the first 12–18 mo after birth, giving rise to a polyclonal population of T cells.

We have demonstrated significant numbers of T cell precursors in the infant intestinal mucosa. Soon after birth, CD4^–8^– (DN) cells accounted for up to 99% of IELs and up to 42% of LPLs. Most of the DN cells expressed CD3 at a moderate or low level, although a CD3^– population was also identified which are likely to be even more immature. CD3⁺ DN cells are likely to be either immature T cells or CD3⁺ NK cells, while DP cells could represent a slightly later stage of immature T cell development (23). However, some DP cells, proposed to be activated T cells, are normally present in adults and are increased in adults with various autoimmune disorders (24, 25). We have observed that a proportion of the DP cells were CD25^– or CD25^low (data not shown), in contrast to in vitro studies which have demonstrated that activated DP cells display an unusually high level of CD25 (26). Thus, it is more likely that the cells we have observed are immature DP cells than activated circulating DP cells. To confirm the likely T cell lineage of CD3^low/– cells, expression of CD2, CD5, and CD7 was studied. CD2 is expressed on a large proportion of developing T cells, before CD3 expression, and by NK cells. CD5 is expressed by T and B cells and their bone marrow-derived precursors. CD7 is a marker of early lineage T cells, expressed before CD3 on nonmyeloid and nonerythroid bone marrow emigrant cells. CD7 may be coexpressed with CD5 before CD5 down-regulation occurs, and can also be expressed by NK cells. Flow cytometry studies revealed that CD7⁺2^–3^– cells accounted for up to 18% of all IEL and 10% of LPL. In contrast, CD2⁺ CD5⁺ cells represented up to 6% of the CD3^– LPL fraction. This combination of cell surface molecules strongly suggests that the cells are immature T cells, supported by immunohistochemistry showing coexpression of CD5 and CD7, a proportion of which were negative for NK, B, and mature T cell markers.

Howie et al. (10) reported CD7⁺ cells in the human fetal intestinal mucosa. Many were CD2^–, especially in the lamina propria (14%, ±7%). They also found a significant proportion (30%) of fetal LPL were CD3⁺CD4^–CD8^–. As in our study, they found that, in the lamina propria, CD4⁺ T cells were more common than CD8⁺ T cells. They did not, however, report CD4⁺ IEL, although we found CD4⁺ cells in the epithelium by flow cytometric and immunohistolochemical analyses. Together, these data suggest that DN T cell precursors accumulate in the mucosa during fetal life and do not differentiate further until after birth, whereupon they gradually mature in situ to SP cells and are joined in the mucosa by thymic emigrants. Evidence of pre-T cells in other tissues of human neonates is limited. Significant numbers of CD34⁺ hemopoietic stem cells are present in both cord and peripheral blood at birth, but decline rapidly with age (27, 28, 29). Hemopoietic cells are present in neonatal liver (30), but evidence for hemopoiesis in the adult human liver is controversial (31, 32).

Immature T cells are likely to be undergoing TCR rearrangement. The RAG protein binds to a pair of recombination signal sequences that flank the V, D, and J gene segments of the germline and initiates a double-strand break that facilitates recombination (33). In mice, intestinal RAG expression has been recognized for a decade (1, 4), although it is usually ascribed to CD8 cells. RAG expression has been reported in human intestine (12, 34, 35), specifically by immature CD3^–CD2⁺TCR^– IEL and CD3⁺TCR^– IEL (34) in the jejunum, and by immature CD3^–CD2⁺CD7⁺ IEL and LPL in the small intestines of adults and children (35). It was recently reported that T cells in the human small intestine and bone marrow express two specific splice variants of the RAG-1 gene (1A/2 and 1A/1B/2) not found in thymocytes (35). It was proposed that T cells using the RAG-1 1A/2 and 1A/1B/2 variants migrate directly from the bone marrow to the small intestine, where they use the 1A/2- and 1A/1B/2-specific RAG-1 variants during TCR gene recombination in situ (35). As our RAG-1 primers are located in exon 2, we would detect all isoforms of RAG-1, and so our finding that 15 of 19 intestinal samples express RAG-1 could correspond to the expression of a gut-specific RAG isoform and certainly merits investigation. After expression of a pre-TCR, consisting of an inframe TCR -chain and pre-T, RAG expression is down-regulated and this is accompanied by a proliferative phase at the end of which CD4 and CD8 are both expressed (DP). After this proliferative phase, RAG is re-expressed and the TCR- gene locus is rearranged. Thus, RAG may be present in both DN and DP T cells. Furthermore, we have demonstrated CD3⁺ DN and DP cells in the human postnatal intestine.

The TEA promoter is located immediately upstream of the TCR-J cluster within the TCR locus (36, 37). It is expressed in immature T cells at the point when DN thymocytes become immature SPs (38), and has been reported in human fetal intestine, but has not previously been investigated postnatally (9, 10). TdT is also present in immature lymphocytes, although the precise stage of T cell maturation has not been defined. Expression of TdT was absent from the intestine of human adults (15) but has not been previously investigated in neonates. Together RAG, TdT, and TEA are associated with ongoing TCR rearrangement and junctional modification. Our finding of these three genes in human intestinal mucosa during infancy strongly supports the hypothesis that T cells are undergoing maturation in the gut mucosa after birth.

We have shown that in early childhood, the TCR T cell repertoire in the small and large intestine is essentially polyclonal. Mucosal, LPL, and IEL cDNA was used as the template for PCR. When TCRBV6 and TCRB12 were studied, very few clones were identified more than once. The population was so polyclonal that, within each individual, few clones were shared between the mucosal sample and either the IEL or LPL fractions. This confirms findings reported by Holtmeier et al. (20), that humans displayed a polyclonal repertoire in midgestation which was retained after delivery, but became oligoclonal during childhood. This is in stark contrast to the findings in adults (18, 19, 39), which have consistently been that the adult intestinal TCR repertoire, at least of IEL, is oligoclonal. In our study only a few samples differed from a polyclonal TCR repertoire. One sample displayed an oligoclonal TCRBV6 repertoire at almost 5 years old, thus being more like the adult repertoire than the other infant samples. By comparing mucosal samples from one patient at 6 and 9.3 mo old, and comparing fractionated LPL in two patients aged 6 and 9 mo old, some restriction of the TCRBV4 repertoire with increasing age seemed to be evident, although in the older mucosal samples from patients aged 12 mo (38% unique) and 18 mo (73% unique) this pattern of increasingly restricted TCRBV4 repertoire was variable.

Together with the T cell phenotype data, then, our findings suggest that, at birth, the intestine is seeded with a polyclonal population of immature T cells. Although relatively few cells were found in the intestine, most of the sequenced TCRB transcripts were different, which implies that they have not undergone clonal expansion. Our data hold true for mucosal samples as well as LPL and IEL fractions, indicating that our findings are not due to experimental artifact. In fact, in experiments not reported here, we have found that mucosal samples of adult intestine show an oligoclonal pattern, which has been widely reported (18, 19, 20). It is possible that either the infant polyclonality is diluted by clonal deletion in situ or migration of clones to other tissues or, more likely, that as the commensal flora stabilizes in later infancy, a few flora-associated clones are favored and expand to become dominant. Changes in chemokine receptor expression during infancy may also contribute to tissue-specific redistribution of clones. In mice, CCL25 is up-regulated upon Ag activation and demonstrates a very restricted pattern of expression essentially limited to the intestine and thymus (40, 41, 42, 43, 44).

TCR junctional length has been the topic of some debate. Moss et al. (45) reported that the length of the junctional region was no longer in adult peripheral blood than in fetal cord blood. However, Hall et al. (46) reported that the proportion of junctional regions expressing 10 or more amino acids was significantly lower in cord blood than in adults. It has been suggested that with greater exposure to Ags, TdT is used to add more nucleotides to increase diversity. Our data suggest that the mean junctional length does not change significantly between birth and 1 year of age. Although TdT transcripts are present and protein is expressed, there is no evidence to support the hypothesis that later rearranged junctional segments have a greater length in response to Ag exposure over time.

The phenotype and repertoire of T cells in intestinal mucosa differs between infancy and adulthood. We have provided evidence for a population of T cells in the infant gut which is immature and undergoing receptor gene recombination amid a polyclonal T cell population. This is taking place at a time of great quantitative and qualitative change in Ag exposure, food and microbial, and it seems likely that this environmental influence must, in some way, shape the repertoire. We were surprised, therefore, that in the two individuals tested in which the intestine had been isolated from environmental influence, there was no difference in clonality from tissues exposed in the normal way. It is possible, then, that these early waves of TCR rearrangement are influenced by absorbed Ags recirculating from other areas of the intestine as well as by locally absorbed luminal Ag. The critical events which cause clonal deletion or selective expansion in late infancy are probably more influenced by microflora and diet, and it is probably these influences which shape the oligoclonal repertoire which is crucial to preventing hypersensitivity and inflammatory responses in the adult gut.

	Acknowledgments

 
We thank Eleri Cusick and Lucinda Huskisson of the Department of Pediatric Surgery for their valued assistance in collecting suitable surgical specimens; and Daniel Carroll and Daniela Vieten for their assistance in obtaining informed parental consent.

	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 funded by the Biotechnology and Biological Sciences Research Council (BBSRC) U.K., Grant No. 7/D13513.

² Address correspondence and reprint requests to Dr. Amanda M. Williams, University Department of Clinical Science at South Bristol, University of Bristol, Old Medicine Building, Bristol Royal Infirmary, Marlborough Street, Bristol, BS2 8HW U.K. E-mail address: Amanda.Williams{at}bristol.ac.uk

³ Abbreviations used in this paper: IEL, intraepithelial lymphocyte; TEA, T early ; DN, double negative; LPL, lamina propria lymphocyte; TCRBV, TCR -chain V region; DP, double positive. SP, single positive.

Received for publication May 7, 2004. Accepted for publication October 4, 2004.

	References

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