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Children’s Research Institute, Departments of
Pediatrics,
Molecular Immunology, Virology, and Human Genetics, and
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Internal Medicine,
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Comprehensive Cancer Center, Ohio State University, Columbus, OH 43205
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Studies in rodents demonstrate that VPAC1 and VPAC2 genes are differentially distributed and regulated (3, 7, 8, 9). Murine VPAC1 is expressed on stimulated and unstimulated thymocytes and splenic CD4+ and CD8+ T cells. In contrast, VPAC2 is expressed on splenic T cell subpopulations only following stimulation. It is expressed at higher levels on splenic CD4+ T cells as compared with CD8+ T cells. Resting and LPS-stimulated murine B cells do not express either VIP receptor (3). VIP binding sites have been detected in rat and mouse peritoneal macrophages (10, 11, 12).
In humans, early studies from our laboratory (13, 14, 15) and by others (16, 17) identified functional high affinity binding sites for VIP in primary T cells, lymphoblastoid cell lines and PBMC. More recently, evidence from immunohistochemistry studies with 125I-labeled VIP have shown a distinct pattern for VIP binding sites. They are colocalized predominantly in T cell areas of human lymph nodes, Peyer’s patches, spleen, and in the cortex and medulla of the thymus (18). However, which receptor subtype is expressed and how these receptors are regulated in different compartments of the human immune system is unknown. The lack of knowledge of the exact distribution and regulation of VIP receptor expression within the human immune system has limited our understanding of the mechanisms of VIP immunomodulation.
Consistent with a differential pattern of expression of VIP receptors in rodents, we hypothesized that VPAC1 and VPAC2 would also have a differential distribution and regulation in human T cell subpopulations and monocytes. The present study used a quantitative fluorescent-based kinetic (real-time) RT-PCR analysis (19, 20) to investigate the VPAC1 and VPAC2 gene expression and regulation within resting and activated human T cell subpopulations (CD4+ and CD8+) and monocytes.
We report the first direct evidence of a differential distribution and selective regulation of VPAC1 and VPAC2 gene expression within peripheral blood T cell subpopulations and monocytes in normal human subjects. We demonstrate a T cell activation-dependent regulation of VPAC1 gene expression and discuss the implications of these findings in understanding the mechanism of VIP as modulator of T cell function.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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This study was approved by the Institutional Review Board of the Children’s Research Institute, Ohio State University (Columbus, OH). Seven healthy volunteers (two females, five males, 21–50 years old) donated blood for isolation of PBMC.
Reagents
Preservative-free mouse mAb anti-human CD3 (clone CRIS-7) and mouse mAb anti-human CD28 (clone 152-2E10) were purchased from BioSource International (Camarillo, CA). EDTA, PMA, and BSA were purchased from Sigma (St. Louis, MO). The stock of PMA was kept at 500 µg/ml in ethanol and the mAb stocks were kept in 100 µg/ml PBS at -20°C. Working dilutions of PMA and mAb were freshly diluted in AIM-V serum-free medium (Life Technologies, Grand Island, NY). EcoRI, HindIII, and XbaI restriction enzymes were purchased from Life Technologies. Aprotinin was purchased from Bayer (Leverkusen, Germany).
T cell and monocyte isolation
PBMC were isolated from EDTA anticoagulated blood by Ficoll-Hypaque gradient (density 1070 ± 0.001 g/ml). (21). Briefly, 100 cc whole blood were diluted 2:1 in PBS, aliquots were layered on 12 ml of Ficoll-Hypaque (Pharmacia, Uppsala, Sweden) in 50-ml tubes in a blood/Ficoll-Hypaque ratio of 3:1 and centrifuged at 400 g for 45 min. The milky interface was recovered, washed twice, and resuspended in PBS with 0.5% BSA and 2 mM EDTA (PBS-BSA-EDTA) for subsequent cell fractionation. Cells for culture were resuspended in AIM-V serum-free medium.
Monocytes and T cell subpopulations were separated in fractions by high gradient magnetic sorting using the Mini and Midi magnetic columns (Miltenyi Biotec, Auburn, CA) as described (22). Briefly, PBMC in PBS-BSA-EDTA, were incubated with anti-CD14 microbeads to positively select monocytes. Subsequently, the CD14- fraction was incubated with a Pan-T cell isolation mixture (Miltenyi Biotec) following the manufacturer’s recommendations to deplete non-T cells and to purify untouched T cells. The depletion mixture consisted of hapten-conjugated Abs (CD11b, monocytes; CD16, neutrophils and a subset of NK cells; CD19, B cells; CD36, platelets; and CD56, subset of NK cells). The incubation with the hapten-conjugated Abs was followed by incubation with microbeads coupled to an anti-hapten mAb for the negative selection of untouched T cells in the magnetic columns.
The enriched T cell suspension was subsequently incubated with CD4
microbeads to positively select CD4+ T
cells; the elution fraction constituted untouched
CD8+ T cells. To account for differences between
positive or negative selection, three of the seven CD4-enriched
suspensions were purified by positive selection of
CD4+ T cells, and two of the five CD8 suspensions
were isolated by positive selection of CD8+ T
cells. Cells were resuspended at a concentration of
107/100 µl final volume, with 80
µl/107 cells of PBS-EDTA-BSA and 20
µl/107 cells of the specific Ab or mixture of
Abs conjugated to the microbeads. After 15 min of incubation with
microbeads (6–12°C), cells were resuspended in cold PBS-BSA-EDTA.
Negatively selected cells were eluted from the column attached to the
magnetic device. Positively enriched cell fractions were eluted from
magnetic columns by removal of columns from magnetic device. Purity of
the cell fractions was measured by flow cytometry. Purity of T cell
fractions was 
95% as judged by flow cytometry analysis using
CD3, CD4, and CD8 mAbs. Monocyte purity was 90% using CD14
mAb.
Flow cytometry
For immunofluorescence staining, cell fractions (CD4, CD8, or CD14+ cells), isolated as described above, were incubated in aliquots of 5 x 105 cells in 50 µl PBS containing 3% FCS and 0.01% NaN3 (PBS-FCS), with saturating concentrations of fluorochrome-labeled mAb for 15 min at 4°C. CD4 PE or PE-Cye5 and CD8 FITC Abs were coincubated, for dual color flow cytometry analysis. The samples were washed twice with 300 µl of the same buffer and resuspended in 100 µl PBS-FCS plus 100 µl of 2% paraformaldehyde. Cell fractions were gated on viable cells and samples were analyzed after fixation on a Coulter (FL) Epics Elite fluorescence-activated cell sorter using 488 (FITC, PE, and R613) and 633 (Cy5) excitation wavelengths. Fluorescence was detected at 525 (FITC), 590 (PE), 613 (R613), and 670 (Cy5) nm. Flow cytometry data was analyzed with the Listmode Coulter Elite software.
Monoclonal Abs
mAbs specific for CD3 (T cells), CD4 (Th cells), CD8 (CTLs) were obtained from Coulter and CD14 (monocytes) from PharMingen (San Diego, CA). Each Ab was titered to determine the highest dilution to give an optimal fluorescent signal. Murine isotype-negative controls were used to establish background fluorescence for each fluorochrome.
Binding of fluorescent VIP
Specific binding of VIP to CD4+ and CD8+ cells was performed by FACS analysis using fluorescein-labeled VIP (Fluo-VIP; NEN, Boston, MA) according to the manufacturer’s recommendations with minor modifications. Cells were washed twice in Earle’s buffer (Life Technologies) supplemented with 20 KIU/ml aprotinin and 0.1% BSA (Earle’s-BSA-APR), centrifuged at 300 x g for 10 min, and aliquoted at 0.5 x 106 cells in 100 µl of Earle’s-BSA-APR buffer. Optimal fluo-VIP concentration and incubation time were determined by titrating the fluo-peptide (10–40 nM) at multiple time points (5, 15, 30 and 45 min). Stock fluo-VIP was kept at 2 µM in DMSO. CD4+- and CD8+-enriched T cell fractions were incubated with fluo-VIP (20 nM) for 5 min in the dark in 96-well round-bottom plates at 37°C. Binding at 4°C was undetectable. Nonspecific binding was determined by competitive binding with saturating concentrations (10 µM) of unlabeled VIP (Bachem, Torrance, CA). Binding reactions were stopped by centrifugation of cells at 4°C followed by washing once with cold Earle’s-BSA-APR. Cells were resuspended in 100 µl of Earle’s-BSA-APR and fixed with 1% paraformaldehyde. Analysis was performed within 24 h of fixing, in a Coulter Elite flow cytometer. Gates were set around a viable-fixed lymphocyte population as determined by forward and side scatter measurements. Cells incubated with unlabeled VIP were used to assess autofluorescence of peptide and cells.
Cell lines
Molt-4b lymphoblasts, a CD4+ T cell leukemia cell line, (American Type Culture Collection, Manassas, VA) were grown as suspension culture in RPMI 1640 medium with 15% heat-inactivated FBS supplemented to final concentrations of 4 mM L-glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin. HT-29 colon carcinoma cells (American Type Culture Collection) were cultured as monolayers in RPMI 1640 with 10% heat-inactivated FBS supplemented as described above. All medium components were obtained from Life Technologies. MHH-MED-1 was a cell line generated from a primary central primitive neuroectodermal tumor at the University of Bonn/Germany (Bonn, Germany) (23). This cell line was cultured in high-glucose DMEM (Life Technologies), 10–15% heat-inactivated umbilical cord serum, and 4 mM L-glutamine. Cells were incubated at 37° in 5% CO2. For RNA isolation, adherent cells (HT-29 and MHH-MED-1) were harvested during exponential growth phase (80–90% confluency). Suspension cells (Molt-4b) were harvested at a density of 1 x 106 or 2 x 106 cells/ml in medium (exponential phase). Cell number was determined by a Coulter counter. Cell viability was determined by trypan blue exclusion technique.
Lymphocyte proliferation
Fractionated T cells (CD4+ and CD8+) were cultured in AIM-V at a final concentration of 1 x 106 cells/ml in 96-well round-bottom tissue culture plates (Falcon; BD Biosciences, Franklin Lakes, NJ) with or without 1 µg/ml preservative-free soluble anti-CD3 (anti-sCD3) alone or in addition to 10 ng/ml PMA or anti-CD3 prebound to plates (immobilized anti-CD3 (anti-bCD3)) alone or in addition to 5 µg/ml soluble anti-CD28. The anti-CD3 was bound to plates as described previously, with some modifications (24). Briefly, 0.2 µg anti-CD3 in 25 µl PBS was added to 96-well plates (for a final concentration of 1 µg/ml) and incubated 2 h at 37°C. Plates were washed once with PBS. Cells (2 x 105/well) were added in suspension in AIM-V medium and the different culture conditions. Cultures were incubated for 72 h at 37°C in a humidified atmosphere of 5% CO2 and pulsed with 1 µCi of [3H]thymidine (NEN) for the final 12 h of incubation. Cells were harvested on a 96-well multiple automated cell harvester (Tomtek, Orange, CT) into a Filtermat A (Wallac, Turku, Finland) and [3H]thymidine quantified in an automated MicroBeta Trilux counter (Wallac).
Quantification of IL-2
Human IL-2 was quantified in cell-free supernatants with an enzyme amplified sensitivity immunoassay kit (EASIA; BioSource International) following the manufacturer’s recommendations. Supernatants were measured undiluted in single wells.
Briefly, a blend of mAbs directed against distinct epitopes of IL-2 was used as capture Abs and was precoated on the microtiter plate. Standards and unknown samples were added into the wells, and a second mAb labeled with HRP directed against IL-2 was added at the same time as the samples. After an incubation period of 2 h on a horizontal shaker, plates were washed (three times) to remove unbound Abs, a chromogenic solution of tetrametylbenzidine was added, and they were incubated for 15 min. The reaction was stopped with 1.8 N H2SO4, and the microtiter plate was read in an ELISA automated reader at 450 nm against a reference filter of 650 nm. Results were extrapolated from a standard curve performed with human recombinant cytokine IL-2. IL-2 was measured in U/ml (sensitivity 0.1 U/ml, 1 U = 83.6 pg/ml).
T cell cultures
Freshly fractionated CD4+ T cells were cultured in AIM-V at a final concentration of 1 x 106 cells/ml in 24-well tissue culture plates (Falcon; BD Biosciences) with or without 1 µg/ml preservative-free anti-sCD3 alone or in addition to 10 ng/ml PMA or anti-CD3 prebound to plates (anti-bCD3, 2 µg/well) alone or in addition to 5 µg/ml soluble anti-CD28. The anti-CD3 was bound to plates as described previously, with some modifications (24). Briefly, 2 µg of anti-CD3 in 150 µl of PBS was added to 24-well plates and incubated the same day for 2 h at 37°C. The plates were washed once with PBS, and cells (1 x 106/ml/well) were added in suspension in AIM-V medium and the different culture conditions. Cultures were set up in triplicates. The plates were incubated for 10 h at 37°C in a humid atmosphere of 5% CO2. Supernatants and cells were harvested at the same time points. Supernatants were harvested for IL-2 determination and cells were preserved in TRIzol reagent (Life Technologies) for VIP, VPAC1, and VPAC2 mRNA analysis. Supernatants and lysates were stored at -80 for <1 mo before analysis.
RNA extraction and cDNA synthesis
Total RNA was isolated from freshly harvested or cultured
CD14+ monocytes, CD4+ or
CD8+ T cells to quantify VIP, VPAC1, and VPAC2
mRNA transcripts. Total RNA was isolated by TRIzol reagent (Life
Technologies) method according to the manufacturer’s protocol. For the
isolation of total RNA from a limited number of cells (
2–3 x
106 cells), the RNAaqueous purification columns
(Ambion, Austin, TX) were used as described in the manufacturer’s
protocol.
Total RNA (1–5 µg) isolated from cells was reverse transcribed using the Superscript Preamplification System for first-strand cDNA synthesis (Life Technologies). The reaction was performed in a total of 20 µl final volume following manufacturer’s protocol for reverse transcription using random hexamer primers; 2.5 µl of the reverse transcription reaction mixture was subsequently used as template for real-time PCR. The same cDNA preparation was used in the analysis of multiple target genes.
Real-time RT-PCR
Real-time RT-PCR was performed using the 5'-3' nuclease activity
of the Taq DNA polymerase enzyme. The method is based on the
direct detection of the amplified product by the release of a
fluorescent reporter dye from a specific fluorescent-labeled probe
during the PCR (19). The probe consists of an
oligonucleotide with a 5'-reporter dye 6-carboxyfluorescein (6-FAM) or
VIC and 3'-quencher dye 6-carboxytetramethylrhodamine (Applied
Biosystems, Foster City, CA). VPAC1-, VPAC2-, and VIP-specific primers
and probes were designed using published cDNA sequences (Table I
) and synthesized by Operon Technologies
(Alameda, CA). VPAC1 and VIP primers span intron/exon junctions. No
information about VPAC2 intron/exon junction sequences was available;
the VPAC2 primer and probe set was tested to insure that it did not
amplify genomic DNA. Ribosomal RNA primers and probe were obtained from
previously published sequences (Taqman ribosomal RNA control
reagent protocol; Applied Biosystems). The probes for VPAC1, VPAC2, and
VIP were labeled with 6-FAM and the rRNA probe was labeled with VIC to
analyze target gene and rRNA in the same well. The rRNA was used to
ensure the quality of RNA preparation, to account for efficiency of the
reverse transcription, and to control for any loading variation of the
initial cDNA amount. Each 5' nuclease assay was performed with two
standard curves, one for VIP, VPAC1, or VPAC2 and the other for
rRNA.
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, A and C) as well
as VIP cDNA or rRNA (not shown) were used to construct standard curves
of copy number vs threshold cycle (CT)(Fig. 1, B and D). VIP and rRNA standard curves were
similarly constructed. VPAC1, VPAC2, and VIP were extrapolated from
each curve as copy number and rRNA as picograms. The standard curve for
VPAC1, VPAC2, and rRNA was constructed with six serial 1/10 dilutions
(standard 1–1 x 108 copies for VPAC1 and
VPAC2, and 500 pg for rRNA) and VIP with five 1/10 dilutions (standard
1–0.8 x 106 copies). The copy numbers were
normalized against 100 pg of rRNA. The results were expressed as copy
number per 100 pg of rRNA using the following formula: (copy number of
VPAC1, VPAC2, or VIP x 100)/picograms rRNA of each individual
sample.
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, A and C). The
fluorescence is detected by a charge-coupled device camera, attached to
the Applied Biosystems Prism 7700 Sequence Detector thermocycler
(Applied Biosystems). The charge-coupled device camera measures the
target-specific signal fluorescent emission spectra from 500 to 650 nm
at defined time intervals (i.e., every 7 s). An amplification
curve was constructed by plotting the CT
(x-axis) vs fluorescent intensity expressed in 
Rn of a given sample (y-axis). The fluorescence
signal increases with each amplification cycle and is proportional to
the starting amount of the cDNA template. The larger the starting copy
number of cDNA, the lower the CT values (Figs. 1 and 2).
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Clones
Ribosomal RNA from PBMC was reverse transcribed using random hexamer primer protocol of Superscript preamplification system for first-strand cDNA kit (Life Technologies). A 187-bp fragment of the ribosomal RNA was then amplified from the reverse transcribed RNA using the published sense (5'CGGCTACCACATCCAAGGAA-3') and anti-sense (5'-GCTGGAATTACCGCGGCT-3') primers (Taqman ribosomal RNA control reagent protocol; Applied Biosystems). These were the same primers used for the real-time PCR. After 10 min of hot start at 94°C, PCR was conducted for 35 cycles for using Gold Taq polymerase (Applied Biosystems) with the following conditions: denaturation 30 s at 94°C, annealing 30 s at 60°C, extension 30 s at 72°C, followed by 10 min of extension at 72°C. The amplicon was cloned into TA vector (Invitrogen, Carlsbad, CA) and sequenced subsequently for confirmation. The cloned fragment was released from the TA vector by digestion with EcoRI. The released cloned fragment was gel purified (Qiagen, Valencia, CA), quantitated, and diluted to the concentration to give a 200-pg/µl volume.
A 534-bp human VPAC1 cDNA fragment was amplified from the human colonic
carcinoma cell line HT-29 by RT-PCR using the following sense and
anti-sense primers: 5'-CTTCTGGTCGCCACAGCTATCCTG-3'; 5'
ACTGCTGTCACTCTTCATGATTAC-3'. The 534-bp RT-PCR product was cloned into
TA vector (Invitrogen) and sequence of the insert was verified by
sequencing of the amplified product. This 534-bp human VPAC1 cDNA
fragment was labeled with [
-32P]dCTP by
random primed labeling (26) and used in screening
106 phages from a human lung cDNA library in
gt11 vector (Clontech, Palo Alto, CA). DNA from positive phage
plaques was isolated by proteinase K treatment (26) and
used in Southern blot analysis. cDNA inserts were released by digestion
with EcoRI, analyzed by agarose gel electrophoresis, and
then subcloned into the EcoRI site of pcDNA3 mammalian
expression vector (Invitrogen). The inserts (clones 1 and 2) were
sequenced. Clone 1 contained the entire coding region (nt 56 to nt 1632
according to the published cDNA sequence (4). The cDNA for
VPAC1 (1576 bp) was released from the vector pcDNA3 using
EcoRI enzyme, gel purified (Qiagen), quantitated, and
diluted to the concentration to give 0.4 x
108 copies/µl of volume.
Human VPAC2 cDNA containing vector human VPAC2/pcDNA3.1+ was kindly provided by Dr. Robert Jensen (National Institutes of Health, Bethesda, MD). The VPAC2 insert (1335 bp) was released from the vector using HindIII and XbaI enzymes, gel purified (Qiagen), and used as standard for the quantitation of the VPAC2 message. VPAC2 clone was quantitated and diluted to the concentration of 0.4 x 108 copies/µl of volume.
Vector pSP6-rVIP was kindly provided by Dr. R. H. Goodman (27). The VIP insert (596 bp) was released from the vector by digesting with EcoRI and HindIII enzymes, gel purified, and used as one of the standards. The clone was quantitated and diluted to a concentration of 3 x 105 copies/µl.
Sequences of all clones were verified in an automated DNA sequencing system (Applied Biosystems 373 DNA sequencer).
Statistical analysis
The Mann-Whitney Rank Sum test or Student’s t test
was used for unpaired data as appropriate. Paired data was analyzed
with Wilcoxon signed rank test and the Kruskal-Wallis ANOVA rank test
with Dunn’s method for multiple comparisons. Sigma Stat (Jandel
Scientific/SPSS Science) statistical software was used for all the
statistical analyses. Statistical significance was measured at
p 
0.05
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Early work from this laboratory identified VIP binding sites, and VIP-dependent cAMP production in human T cells (13). To delineate whether VPAC1 or VPAC2 is expressed in monocytes and T cell subpopulations (CD4+ and CD8+), cells were purified and total RNA was extracted for quantitative analysis of VPAC1 and VPAC2 transcripts by kinetic RT-PCR.
Our results indicate that the major receptor transcribed in peripheral
blood resting T cells and monocytes is VPAC1 (Fig. 2, A and
C). Within human PBMC, resting monocytes and
CD4+ T cells have higher levels of VPAC1 relative
to CD8+ T cells (p
0.05). Both VPAC1 and VPAC2 are expressed in resting T cells (Fig. 2, A and C), but VPAC1 transcripts are significantly
higher relative to VPAC2 on CD4+ T cells,
(mean ± SE, 1451 ± 493 vs 62 ± 18 copies/100 pg rRNA;
p < 0.05, n = 7) and on
CD8+ T cells (mean ± SE, 154 ± 51 vs
66 ± 31 copies/100 pg rRNA; p < 0.05,
n = 5). Resting monocytes expressed only VPAC1
(mean ± SE, 2914 ± 940 copies/100 pg rRNA;
n = 7); VPAC2 was below detectable levels (<50
copies/100 pg rRNA; n = 7).
Fig. 2, B and D, demonstrates VPAC1 and
VPAC2 amplification plots in resting CD4+ and
CD8+ T cells with HT-29 cells as a positive
control for VPAC1 expression and Molt-b cells as positive control for
VPAC2.4 VPAC1
transcripts are most abundant in HT29 >
CD4+ > CD8 with no detectable VPAC1 transcripts
in Molt4b cells (Fig. 2B). VPAC2 transcripts are most
abundant in Molt4b > CD4+ =
CD8+ with no detectable VPAC2 transcripts in HT29
cells (Fig. 2D).
No significant differences were observed in VPAC1 or VPAC2 expression levels based on positive or negative selection of CD4+ or CD8+ T cell suspensions. Three of seven CD4+ T cell suspensions and two of five CD8+ T cell suspensions analyzed were positively selected; consequently, results of positively and negatively selected cell populations are combined in this analysis.
Differential expression of VIP receptor protein on resting CD4+ and CD8+ T cells
To test whether differential VPAC1 gene expression in T cell
subpopulations resulted in differential VIP binding in
CD4+ or CD8+ T
cell-enriched suspensions, positively selected
CD4+ (n = 3) or
CD8+ T cells (n = 3) were
incubated with fluo-VIP under optimal binding conditions as described
in methods. Fig. 3A
demonstrates specific binding of fluo-VIP to CD4+
cells as shown by a shift to the right in the fluorescence intensity of
the cells incubated with 20 nM fluo-VIP (total binding) compared with
cells incubated with fluo-VIP plus 10 µM unlabeled VIP (nonspecific
binding). Specific binding of fluo-VIP to CD4+ T
cells varied from 4.5% to 43.3%. CD8+ T cells
have no detectable specific binding of fluo-VIP as shown in Fig. 3B. The background fluorescence for suspensions incubated
with unlabeled VIP alone was <1% in all CD4 and CD8 suspensions
tested (n = 6).
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There are no previous reports on regulation of VPAC1 and VPAC2
expression in human T cells. Fig. 4
demonstrates the effect of T cell activation with PMA alone or in
combination with anti-sCD3 on expression of VPAC1 and VPAC2 mRNA
transcripts.
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A). In contrast, VPAC2
copy number increased in three of the four anti-sCD3 plus
PMA-stimulated cultures, but these changes were not statistically
significant (means ± SE, medium, 290 ± 163 vs anti-sCD3
plus PMA, 556 ± 246 copies/100 pg rRNA; p = 0.18,
n = 4) (Fig. 4B). VIP mRNA was not
detectable (<50 copies/100ng rRNA) in resting, PMA-stimulated, or PMA
plus anti-sCD3-activated CD4+ cells
(n = 3).
CD4+ T cells were activated by PMA and
anti-sCD3 as shown by induction of IL-2 and cell proliferation
(Fig. 4, C and D). Low, but detectable IL-2
levels were observed in resting CD4+ T cells
(0.75 ± 0.14 U/ml, mean ± SEM; n = 3); PMA
alone induced moderate IL-2 (17.1 U/ml; n = 1) while
PMA plus anti-sCD3 induced maximal IL-2 production (69.3 ±
6.29 U/ml; p < 0.01, n = 3). Cell
proliferation was also documented in parallel cultures as measured by
[3H]thymidine uptake (Fig. 4D); PMA
plus anti-sCD3 induced proliferation relative to unstimulated
controls (161.3 ± 28 vs 70685 ± 23673, mean ± SEM;
p < 0.05, n = 3).
Activation by TCR cross-linking and costimulation results in down-regulation of VPAC1 in CD4+ T cells
VPAC1 was down-regulated (71.2% reduction relative to
unstimulated controls, p = 0.12, n = 2)
in anti-bCD3 plus anti-sCD28-stimulated cultures (Table II). VPAC2 was not up-regulated nor was
VIP mRNA detectable in purified stimulated CD4+ T
cells under these experimental conditions, (data not shown).
Cross-linking the TCR/CD3 receptor complex and the CD28 costimulatory
pathway in negatively selected CD4+ cells
resulted in activation as determined by IL-2 production. Increased IL-2
was detected in anti-bCD3 plus anti-CD28 relative to
unstimulated controls (p < 0.05,
n = 2) (Table II). Th cell proliferation was assessed
in parallel cultures by [3H]thymidine
incorporation in anti-bCD3 plus anti-CD28 treated
CD4+ T cells relative to unstimulated controls
(p = 0.078, n = 2).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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There are no previous quantitative analyses of VPAC1 and VPAC2 gene expression in any species. Our results are in agreement with and expand previous studies of VIP receptor expression by conventional RT-PCR in rodents. These studies show that VPAC1 is constitutively expressed and that VPAC2 is inducible in bulk thymocytes and splenocytes and in fractionated CD4+ and CD8+ T cells (8, 9, 28). Our results demonstrate very low but detectable levels of VPAC2 in resting CD4+ and CD8+ T cells (mean, 62 copies and 66 copies/100 pg rRNA, respectively), in contrast with previous studies in rodents (8, 9, 28). This difference may be due to difference in sensitivity between conventional RT-PCR and real-time RT-PCR. The results of our study on VPAC2 up-regulation are similar to those in rodents. VPAC2 in both mouse and human fractionated T cells appear to require two stimuli, anti-sCD3 and PMA.
There are no previous reports of differences between the level of gene expression of VIP receptors in T cell subpopulations (CD4 and CD8). However, predominant VIP binding on CD4 relative to CD8 has been shown previously (17) and our fluo-VIP binding results confirm that observation. We are extending this observation by additionally demonstrating predominant VPAC1 gene expression on resting CD4+ T cells vs CD8+ T cells, which may explain these differences. VPAC2 has very low levels of expression in both cell populations.
There are no previous reports of T cell activation-induced down-regulation of VPAC1 gene expression. Other receptor/ligand systems (neurotensin) have demonstrated that activation of receptor gene transcription is required to maintain cell sensitization after agonist exposure (29). VIP/VPAC1 may be similarly regulated. Our results on activated Th cells, demonstrating fewer VPAC1 transcripts, complement and expand those of Johnston et al. (30) showing less responsiveness to VIP-induced chemotaxis. In the latter study, down-regulation of the number of 125I-labeled VIP binding sites was observed in purified T cells (predominantly in Th cells) as a result of cross-linking the TCR/CD3 receptor complex. We used a similar experimental design in that we cross-linked the TCR/CD3 receptor and stimulated purified Th cells. We observed down-regulation of VPAC1 gene expression and no changes in VPAC2 expression in activated Th cells, clearly demonstrating differential regulation of VPAC1 and VPAC2. Thus, Johnston et al.’s results and our new observations together strongly suggest that T cell activation-induced down-regulation of 125I-labeled VIP binding sites is a result of down-regulation of the VPAC1 gene expression.
Occupancy of the TCR/CD3 receptor by anti-sCD3 induced minor changes in VPAC1 and VPAC2 gene expression. Consistent with previous reports (31, 32), anti-sCD3 was not able to induce proliferation in purified CD4+ and CD8+ T cells in the absence of a second signal. Occupancy of the TCR/CD3 receptor with anti-sCD3 has been associated only with increased intracellular Ca2+ concentration (32), whereas the addition of PMA is associated with activation of PKC (31). Sustained activation of PKC has been shown to be important for subsequent T cell proliferation (33). These differences may account for the lack of effect of anti-sCD3 on VPAC1 in contrast to the effects of anti-sCD3 plus PMA and PMA alone, which suggest a role for PKC in the modulation of gene expression of VPAC1 and VPAC2.
T cell activation with either anti-sCD3 plus PMA or the combination of anti-bCD3 plus anti-CD28 was effective in down regulating VPAC1 gene expression. We observed a gradient effect in the down-regulation of VPAC1 that was inversely proportional to the levels of IL-2 and proliferation. Those in vitro conditions that induced higher levels of IL-2 and induced optimal levels of proliferation, (anti-sCD3 plus PMA and anti-bCD3 plus anti-CD28) also reduced most significantly the VPAC1 gene expression. VIP via VPAC1 is a potent inducer of cAMP (13, 15), and inhibitory effects of high levels of cAMP on T cell proliferative responses are well-documented (34, 35). Whether T cell activators would down-regulate the VPAC1 gene expression before progressing toward optimal IL-2 secretion and proliferation warrants further investigation.
The suppressive effect of PMA on 125I-labeled VIP binding has been demonstrated in early studies of VIP receptor internalization in HT-29 cell lines (36). This study demonstrated that the effect of PMA was mediated via PKC activation and independent of agonist-induced down-regulation of the VIP receptor. Our results confirm and expand this previous study and would suggest that PMA and anti-CD3 may regulate VPAC1 gene expression at the transcriptional level. We speculate that the suppressive effect of PMA and anti-sCD3 on VPAC1 gene expression may be mediated by a cis regulatory repressor element(s) located at the 5'-flanking and promoter region of the VPAC1. Recently, Pei (37) cloned a repressor protein that down-regulates VPAC1 expression in rats; an homologous VPAC1 repressor protein in humans has yet to be identified. Whether PMA and anti-CD3 effects on VPAC1 may be mediated by this mechanism is as yet untested.
PMA induced suboptimal IL-2 synthesis and suboptimal proliferation on purified CD4+ T cells under our experimental in vitro conditions. Earlier studies have documented that c-myc and c-fos transcriptional activation can be induced by an increase in either intracellular Ca2+ concentration or activation of PKC alone (38), which could explain the suboptimal T cell activation by PMA alone. The effect of PMA on VPAC1 down-regulation also suggests a partial effect of PKC activation on the transcriptional regulation of these genes that is synergistic with anti-sCD3. The synergistic effect of the combined stimuli (PMA plus anti-sCD3) is well documented in T cell activation and is shown here when anti-sCD3 plus PMA induced optimal IL-2 synthesis and proliferation of the CD4+ T cell suspensions. Taken together, the results regarding activated CD4+ T cells suggest that a decrease of VPAC1 gene expression precedes optimal proliferation of T cells activated via anti-sCD3 plus PMA or bound anti-CD3 plus anti-CD28.
VIP has been shown to have anti-proliferative effects on T cells by decreasing IL-2 synthesis associated with VIP-induced cAMP (40). Because VPAC1 is a significantly more potent inducer of cAMP than VPAC2, VPAC1 likely mediates the inhibitory effects of VIP.4 One potential implication of the reported selective down-regulation of VPAC1 in activated cells could be that VIP might only exert its inhibitory effect on resting T cells that have persistent VPAC1 expression. This may be particularly important in the context of selective activation of T cells (i.e., antigen-specific response). Thus, VIP may play a role in limiting bystander activation of T cells, which is an important aspect of the regulation of the immune response.
Resting monocytes express VPAC1 exclusively. In contrast to our observations, Dewit et al. (41) report VPAC2 expression in human resting enriched monocyte suspensions. These differences are probably due to different methods of isolation (positive selection by magnetic beads conjugated CD14 vs plate-adherence method). Because VPAC2 is expressed at very low levels in resting T cells but is up-regulated in activated T cells, contamination of monocyte preparations with activated T cells could be a source of VPAC2. Alternatively, the isolation procedure, that includes an incubation in serum supplemented medium at 37°C may induce VPAC2 expression in the monocytes. Similar to our findings, resting peritoneal macrophages of the mouse and the rat also constitutively express VPAC1 (10, 42, 43) and not VPAC2.
The source of VIP that lymphocytes encounter has been controversial, (reviewed in Ref. 44). Our current results are in agreement with previous results from this laboratory (45) in which neither resting nor activated CD4+ or CD8+ T cells and monocyte suspensions analyzed expressed detectable levels of VIP transcripts. Thus, the role of VIP as an autocrine or paracrine mediator may be questionable. We favor the view that the primary source of VIP that interacts with T cells or monocytes is the VIPergic nerves in the microenvironment of the T cells in lymph nodes, Peyer’s patches, thymus, and spleen (46).
The results presented in this study suggest that selective expression and differential regulation of VPAC1 and VPAC2 may be one of the mechanisms of control of VIP modulation of T cell and monocyte function.
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2 Address correspondence and reprint requests to Dr. M. Sue O’Dorisio, University of Iowa, Division of Pediatric Hem/Oncology, 2524 JCP, 200 Hawkins Drive, Iowa City, IA 52242.
3 Abbreviations used in this paper: VIP, vasoactive intestinal peptide; VPAC1, VIP receptor type 1; VPAC2, VIP receptor type 2; anti-sCD3, soluble anti-CD3; anti-bCD3, immobilized anti-CD3; Earle’s-BSA-APR, Earle’s buffer with 20 KIU/ml aprotinin and 0.1% BSA; Fluo-VIP, fluorescein-labeled VIP; CT, threshold cycle; 6-FAM, 6-carboxyfluorescein.
4 M. A. Summers, M. S. O’Dorisio, M. L. Lara-Marquez, S. P. Sreedharan, and E. J. Goetzl. Identification of a fragment of vasoactive intestinal peptide (VIP4–28) as an endogenous antagonists for VIP receptor type 2 (VPAC2). Submitted for publication.
Received for publication September 22, 1999. Accepted for publication December 5, 2000.
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