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Prepublished online as a Blood First Edition Paper on January 23, 2003; DOI 10.1182/blood-2002-08-2671.
TRANSPLANTATION
From the Fred Hutchinson Cancer Research Center
(FHCRC), Seattle, WA, and University of Washington, Seattle.
In mice, interleukin-7 (IL-7) hastens T-cell reconstitution and
might cause autoimmune diseases, lymphoma, and osteoporosis. We
assessed the effect of IL-7 on T-cell reconstitution and toxicity in
baboons that underwent total body irradiation followed by autologous transplantation of marrow CD34 cells. Three baboons received placebo and 3 baboons received recombinant human IL-7 (rhIL-7, 75 µg/kg twice
a day subcutaneously) between 6 and 10 weeks after
transplantation. The mean increase in blood absolute CD4 T-cell counts
was 0.9-fold in the placebo-treated animals versus 9.0-fold in those
treated with IL-7 (P = .02). The increase observed in the
IL-7-treated animals appeared attributable to peripheral expansion
rather than de novo generation. The IL-7-treated animals had greater
mean increases in the volumes of the spleen (2.0-fold with placebo versus 4.5-fold with IL-7, P = .02) and lymph nodes
(1.8-fold with placebo versus 4.1-fold with IL-7,
P = .10) but not the thymus (3.4-fold with placebo versus
1.1-fold with IL-7, P = .18). Side effects of IL-7
included thrombocytopenia and possibly neutropenia and hemolytic
anemia. One IL-7-treated animal failed to thrive due to a disease
resembling graft-versus-host disease. No animals developed lymphoma.
Bone density was not decreased. In conclusion, IL-7 raises CD4 T-cell
counts in irradiated primates. It remains to be determined whether this
is associated with clinical benefit.
(Blood. 2003;101:4209-4218) A reduction in the number of T cells results in a
homeostatic attempt to increase the T-cell number.1-9 This
occurs either through the differentiation of marrow-derived progenitors
into T cells in the thymus (de novo generation) or through the
proliferation of existing T cells (peripheral
expansion).10-12 In mice, interleukin 7 (IL-7) is a potent
stimulator of both de novo generation and peripheral
expansion.13 Peripheral expansion appears stimulated through stimulation of proliferation and inhibition of cell
death.14-16 In patients with HIV or chemotherapy-induced
lymphocytopenia, there is an inverse correlation between CD4 T-cell
counts and IL-7 levels.17,18 Thus, IL-7 may play a
significant role in T-cell homeostasis.
After hematopoietic cell transplantation, patients have frequent
infections for at least 1 year. This is in part because CD4 T-cell
reconstitution proceeds slowly (over months to years; for reviews, see
Storek and Witherspoon3 and Parkman and
Weinberg19). Peripheral expansion may be limited because T
cells from transplant recipients are prone to apoptosis.20
De novo generation may be reduced from conditioning-induced damage to
thymic stromal cells, which secrete IL-7.21 IL-7 levels
after transplantation are only mildly elevated
(~ 2-fold22) possibly due to the damage to thymic
stroma. Administration of IL-7 to mice after transplantation improved
both de novo generation and peripheral expansion of CD4 T
cells.23-25 These appeared to be of clinical benefit
because the mortality rate after pulmonary infection with influenza
virus in mice receiving IL-7 was 44% versus more than 90% in mice not receiving IL-7.24
B-cell reconstitution was also hastened in IL-7-treated mice receiving
transplants.23 This could be of added benefit because B
lymphocytopenia appears to contribute to the development of infections
in transplant recipients.26
Before starting human trials we wished to evaluate IL-7 in
nonhuman primates, for 3 reasons. First, the function of IL-7 in rodents and primates may be different.27-30 It is
conceivable that IL-7 could hasten T- or B-cell reconstitution in
rodents but not in primates. Second, in nonhuman primates one can
assess T and B cells not only from blood but also from tissues. The
tissue lymphocytes constitute the vast majority of total body
lymphocytes and may differ from circulating lymphocytes; changes in
their quantity are not always reflected by parallel changes in their blood count.8,31-35 Third, we wished to determine the side
effects of IL-7 in nonhuman primates, which should significantly
influence the enthusiasm for and the design of clinical trials. In this study, the following potential side effects were evaluated particularly carefully: (1) lymphoma/thymoma because IL-7-transgenic mice developed lymphomas and thymomas36,37; (2) autoimmunity because
IL-7-transgenic mice developed ulcerative colitis or dermal T-cell
infiltrate with alopecia, hyperkeratosis, and
exfoliation37,38; and (3) osteoporosis because healthy
female mice treated with IL-7 for 20 days developed osteoporosis of the
same degree as estrogen-deficient (oophorectomized) mice, due to
increased osteoclastic resorption.39
Study design and animals
Transplantation and posttransplantation treatment
Escherichia coli-produced rhIL-7 (kindly provided by Dr Michel Morre, Cytheris, Vanves, France) was administered subcutaneously for 28 days. The IL-7 was diluted in 20 mM sodium citrate buffer, pH 6.2. The same buffer was used as placebo. Enumeration of MNC subsets Mononuclear cells (MNCs) were separated from EDTA (ethylenediaminetetraacetic acid)--anticoagulated blood using density gradient (Ficoll, density 1.073 kg/L) centrifugation. The cells, suspended in flow buffer (phosphate-buffered saline [PBS] with 1% bovine serum albumin and 0.1% sodium azide) were stained with fluorochrome-conjugated monoclonal antibodies and analyzed by 3-color flow cytometry. The following antibodies were used: CD3 from Biosource (clone FN18; Camarillo, CA), CD4 from BD Biosciences (clone SK3; San Jose, CA), CD8 from Beckman-Coulter (clone B9.11; Miami, FL), CD11a from Beckman-Coulter (clone 25.3.1), CD14 from Beckman-Coulter (clone RMO52), CD16 from Beckman-Coulter (clone 3G8), CD20 from BD Biosciences (clone L27), CD45RA from Beckman-Coulter (clone ALB11), and CD56 from Beckman-Coulter (clone N901). Residual red blood cells were lysed in hemolytic buffer (8.3% ammonium chloride plus 1% potassium bicarbonate plus 0.4% EDTA). No fixation was done. Cells were resuspended in flow buffer and kept on ice until flow cytometry, which occurred within 12 hours from blood drawing. To identify apoptotic cells, cells were stained not only with the fluorochrome-labeled antibodies against surface antigens, but also with either fluorochrome-labeled annexin V (using 2.5 mM CaCl2 in 140 mM NaCl/10 mM HEPES [N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid]/NaOH, pH 7.4) or 7-actinomycin D (7-AAD, 20 mg/L; Sigma, St Louis, MO).FACSCalibur (BD Biosciences) flow cytometer was used for data acquisition. List-mode data were analyzed using WinList software (Verity Software House, Topsham, ME). The forward scatter × side scatter (FSc × SSc) gate was set to encompass MNCs. B cells were defined as CD20+ MNCs. CD4 T cells were defined as CD3+CD4+ MNCs, and CD8 T cells were defined as CD3+CD8+ MNCs. Natural killer (NK) cells were defined as MNCs expressing CD16 or CD56 and not expressing CD3 or CD14. Monocytes were defined as CD14+ MNCs. Each absolute MNC subset count was calculated as the absolute MNC count multiplied by the percent of the MNC subset divided by 100. The absolute MNC count represented the sum of absolute lymphocyte count and absolute monocyte count determined by our clinical hematology laboratory, using Sysmex XE2100 cell counter and Giemsa-stained blood smears. Lymph nodes (LNs) were obtained at 6 weeks (before starting IL-7, the largest LN from the right axilla), 10 weeks (approximately 12 hours after the last injection of IL-7 or placebo, the largest LN from the left axilla), and approximately 6 months after transplantation (the largest LN from the right axilla). Axillary LNs were chosen because they were distant from the IL-7/placebo injection sites (thighs). The LNs were teased in RPMI with 10% fetal calf serum. The resulting cell suspension was filtered through a 70-µm nylon cell strainer. Percentages of cell subsets were determined by flow cytometry as in the blood. TREC assay First, we determined the nucleotide sequence of the signal joint of baboon  T-cell receptor excision circle (TREC). In rhesus macaques and sooty mangabeys the signal joint sequence was found to be
approximately 96% homologous to that in humans.35 Thus, we hypothesized that the baboon sequence would also be similar. Polymerase chain reaction (PCR) amplification of the putative signal
joint from baboon blood MNC DNA, using primers for regions 100%
homologous among humans, macaques, and mangabeys (forward primer,
5'-CACATCCCTTTCAACCATGCT-3'; reverse primer,
5'-TTGCTCCGTGGTCTGTGCTGGCATC-3'), yielded a product of the expected
length (281 bp). Sequencing of the product showed that baboon and human
signal joint regions are 94.4% homologous (GenBank accession no.
AF541878). The 281-bp PCR product represented a TREC because
it was detected in DNA extracted from fluorescence-activated
cell-sorted (FACS) normal baboon CD45RAhigh CD4 T cells but
not CD45RA CD4 T cells or B cells.
TREC levels were determined by real-time PCR, analogous to Douek et al.42 FACS CD4 or CD8 T cells were lysed in 100 µg/mL proteinase K at 107 cells/mL. The 5'-nuclease (TaqMan) assay was performed on 5 µL cell lysate (total volume of the lysate of 50 000 cells was 40 µL; thus 5 µL lysate contained TRECs from 6250 cells), using forward primer, 5'-CACATCCCTTTCAACCATGCT-3'; reverse primer, 5'-GCCAGCTGCAGGGTTTAGG-3'; and probe FAM-ACGCATTTGGTTTTTGTAAAGGTGCTCACT-TAMRA (MegaBases, Chicago, IL). PCR reactions contained 0.5 U Taq polymerase, 3.5 mM MgCl2, 0.2 mM dNTPs, 400 nM each primer, 200 nM probe, and Blue-636 reference (MegaBases). The reactions were run at 95°C for 5 minutes, then 95°C for 30 seconds and 60°C for 1 minute for 40 cycles, using ABI7700 system (PE Biosystems, Norwalk, CT). Samples were analyzed in quadruplicates. Plasmids containing the baboon signal joint region were used as standards. A standard curve was plotted and the TREC level (the number of TREC copies per 6250 CD4 or CD8 T cells) was calculated using the ABI7700 software. TREC levels reflect not only the de novo generation of T cells but also the expansion of thymocytes or T cells after rearrangement. For example, TREC levels drop when peripheral T cells proliferate as a result of an infection. In contrast, the absolute count of TREC-containing (TREC+) T cells (per unit blood volume) is not influenced by rearrangement after proliferation and thus serves as a better indicator of the quantity of T cells generated de novo. We calculated the absolute count of TREC+ CD4 (CD8) T cells (per microliter) as the TREC level (per 6250 cells) multiplied by the absolute CD4 (CD8) T-cell count (per microliter) and divided by 6250. CT For oral contrast, 0.8% barium sulfate was used (E-Z-CAT, E-Z-EM, Westbury, NY). The scanning was done on GE Lightspeed QX/I computed tomography (CT) scanner, using the following conditions: abdominal protocol, 120 kV, 130 mA, 1.25-mm slice thickness, 21.3-cm diameter of field of view, and 102-cm table height. After scanning without intravenous contrast, 68% ioversol (Optiray 320, Mallinckrodt, St Louis, MO) was injected intravenously from a power injector (1 mL/s, total of 48 mL). Using the same conditions as for the nonintravenous contrast scanning, scanning with intravenous contrast was started 1 minute after finishing the intravenous injection. On analysis, the area of the thymus was determined on each slice (mm2), using intravenous contrast images and GE Pathspeed software (GE Medical Systems, Milwaukee, WI). The area was multiplied by the slice thickness (1.25 mm) to get the volume at that slice level (mm3). Thymic volume was calculated as the sum of the volumes at each slice level. Splenic and LN volumes were determined analogously. The sum of the volumes of one right axillary LN, one left axillary LN, one pelvic LN, one right inguinal LN, and one left inguinal LN was used as an index of the size of all lymph nodes. In each LN region, the largest LN at week 6 that was also identifiable at weeks 10 and 14 was measured.Bone density was measured using a calibration phantom (Image Analysis, Columbia, KY) placed under the lumbar area of the animal at the time of scanning. Nonintravenous contrast images and GE PathSpeed software were used for analysis. On each slice of interest, the mean Hounsfield unit (HU) of the vertebral body cancellous bone and the mean HU of each of the 4 density standards were determined. Density of the vertebral body cancellous bone was calculated by linear regression. This was done at 3 levels of vertebra L3 (1.25 mm above the basivertebral canal, at the basivertebral canal, and 1.25 mm below the basivertebral canal), 3 analogous levels of vertebra L4, and 3 analogous levels of vertebra L5. For each vertebra, the mean density of the 3 levels was calculated. The mean density of the mean densities of L3, L4, and L5 ("L3-5 density") was used as the index of overall cancellous bone density. We focused on cancellous bone because in states of increased osteoclastic resorption the density of cancellous bone is decreased to a greater degree than the density of solid bone. For quality control, a torso phantom (Image Analysis) was used with each scanning. Its calculated density ranged from 96.1 to 105.9 mg/cm3, suggesting adequate reproducibility. Histology Using standard techniques, buffered formalin-fixed, paraffin-embedded and, in the case of bone marrow, decalcified tissue sections were stained with hematoxylin and eosin (H and E) or immunostained with CD3 (rabbit-antihuman CD3 polyclonal antibody, Dako, Carpinteria, CA), CD4 (clone 1F6, Novocastra, Newcastle upon Tyne, United Kingdom), or CD20 (clone L26, Dako). Snap-frozen tissues were immunostained with CD8 (clone G10.1, kind gift from Dr Bing Hu, University of Washington, Seattle). Slides were analyzed by one pathologist (R.C.H.) blinded to IL-7 versus placebo treatment. Dermal perivascular T-cell infiltrates were given a score from 0 to 3. Bone marrow cellularity was estimated from the sections stained with H and E and expressed as the percentage of nonfat cells among fat plus nonfat cells. Percentage of T or B cells among bone marrow hematopoietic cells was estimated from the immunostained sections.IL-7 levels and neutralizing antibodies Serum concentration of IL-7 was determined by enzyme-linked immunosorbent assay (ELISA) using the kit for human IL-7 (R & D Systems, Minneapolis, MN). In case of values exceeding the concentration of the highest standard, diluted serum was used.For the detection of IL-7-neutralizing antibodies, IL-7-dependent
cells (PB1, from DSMZ [German Tissue Culture Collection]) were grown
in 85% McCoy 5A medium supplemented with 15% fetal bovine serum
(Hyclone, Logan, UT), 50 ng/mL human IL-7 (Cytheris, Vanves,
France), 0.4 × minimum essential medium (MEM) vitamins (Sigma), 1 × MEM nonessential amino acids (Sigma), 0.5 × essential amino acids (Sigma), 1 mM sodium pyruvate, 0.12% sodium bicarbonate, 2 mM L-glutamine, and 50 µM Statistics Increases in MNC subset counts were calculated as week 10 count/week 6 count. The t test was used to test differences in the increases between the IL-7 and the placebo-treated animals (SigmaStat software, SPSS, San Rafael, CA). Two-tailed P values are given.
Baboon and human IL-7 are similar The mRNA from the thymus of a normal baboon was extracted and reverse transcribed. The cDNA was PCR amplified, using primers for the flanking regions of the human IL-7 open reading frame (forward, ATGTTCCATGTTTCTTTTAGGTA; reverse, TGATGGGCACTAAAGAACACTGA), and sequenced. The resulting baboon IL-7 open reading frame (GenBank accession no. AF541946) was found to have 98.1% nucleotide sequence homology and 96.6% amino acid sequence homology with human IL-7.43 Given the high degree of homology between baboon and human IL-7 and because human IL-7 has been shown to stimulate T-cell reconstitution in mice,23-25 whose IL-7 amino acid sequence is only about 80% homologous with that of humans,43,44 we hypothesized that human IL-7 should have biologic activity in baboons.Effect of IL-7 on CD4 T cells In the main experiment involving 3 IL-7-treated and 3 placebo-treated animals, CD4 T-cell counts in the blood increased from week 6 to week 10 on average 9.0-fold in the IL-7-treated compared to 0.9-fold in the placebo-treated animals (P = .02; Figure 1A). CD4 T-cell counts also increased in the 2 animals in the pilot experiment (J99202, 5.7-fold; A00066, 1.9-fold).
The differences between IL-7- and placebo-treated animals at week 10 appeared to be greater for memory/effector (CD45RAlow/
It is unlikely that the increase in circulating CD4 T-cell counts is
due to an IL-7-stimulated shift of CD4 T cells from tissues to blood.
On the contrary, there was evidence of increased T-cell counts in
tissues sampled (LNs, bone marrow, and skin). Between week 6 and 10, LN
volumes increased more in the IL-7- than placebo-treated animals (mean
4.2- versus 1.8-fold increase; Figure 2), whereas the changes in the
percentages of CD4 T cells among LN cells were similar (mean 0.6- versus 0.6-fold increase; Figure 3),
indicating that total LN CD4 T cells increased more in IL-7- than
placebo-treated animals. Marrow at week 10 tended to be more cellular
in IL-7- than placebo-treated animals (mean 99% versus 88%,
P = .18) and was heavily infiltrated with T cells (mean
75% versus 9% T cells among hematopoietic cells,
P < .001; Figure 4).
Likewise, the skin on week 10 tended to contain more T cells in IL-7-
than placebo-treated animals, particularly around dermal capillaries,
both at the IL-7/placebo injection site (thigh, mean perivascular
T-cell score 2.7 versus 2.0, P = .12) as well as at the
noninjection site (arm, mean perivascular T-cell score 2.7 versus 1.5, P = .12). Collectively, these results suggest that IL-7
treatment resulted in increased total body CD4 T-cell counts.
The IL-7-stimulated peripheral expansion leading to the increased
total body CD4 T cells could result from increased proliferation or
decreased death rate or both. IL-7- and placebo-treated animals had
similar week 6 to 10 changes in the percentages of annexin V+ or 7-AAD+ CD4 T cells (Figure
5). This suggests increased proliferation as the mechanism of the IL-7-stimulated peripheral expansion of CD4 T
cells. However, a direct determination of the proliferation (eg, using
Ki67) is needed to confirm this hypothesis.
Effect of IL-7 on CD8 T cells, B cells, NK cells, and monocytes Compared with CD4 T cells, other MNCs were affected by IL-7 to a lesser degree or not at all. IL-7 treatment seemed to be associated with a mild increase in CD8 T cells (Figure 6). This was also consistent with peripheral expansion rather than de novo generation because CD11ahigh (memory/effector) CD8 T cells and not CD11alow (naive) or TREC+ CD8 T cells increased more in the IL-7- than placebo-treated animals (data not shown). IL-7 did not increase counts of B cells in blood or LNs; on the contrary, IL-7 may have mildly delayed B-cell reconstitution (Figure 6). Consistent with this, marrow and skin B cells were undetectable or barely detectable at week 10 in both IL-7- and placebo-treated animals. Monocyte and NK cell counts were unaffected (data not shown).
Potential toxicity of IL-7 Platelet counts were lower in IL-7-treated than placebo-treated animals (Figure 7). This may have resulted from decreased marrow production because megakaryocytes at week 10 appeared to be decreased in IL-7 compared with placebo-treated animals (Figure 4). Decreasing platelet counts during IL-7 treatment from more than 100 to less than 40 × 109/L were also noted in J99202 and A00066 (the pilot experiment animals that received IL-7 after 3 months after transplantation).
Severe neutropenia ( Hemolytic anemia developed in 2 of 3 IL-7-treated animals (K99307 and M99267) and 0 of 3 placebo-treated animals. In both cases, nadir hemoglobin levels were below 7 mg/dL, peak reticulocyte counts above 140 × 109/L (above 10%), and nadir haptoglobin levels below the detection limit. Blood smears were negative for intraerythrocytic parasites. Direct antiglobulin (Coombs) test was weakly positive for complement in K99307 and negative in M99267; however, a negative test does not rule out autoimmune etiology because the antihuman antibody used in the assay (kit purchased from Ortho Clinical Diagnostics, Raritan, NJ) may not cross-react with baboon antigens. In K99307 the anemia was diagnosed on day 62, was associated with a cold agglutinin, and recovered by day 125, possibly due to the discontinuation of IL-7 on day 69 or warming the animal with a heat lamp. In animal M99267 the anemia was diagnosed on day 153, was not associated with a cold agglutinin, and responded to prednisone. Decreasing hemoglobin levels during IL-7 treatment were also noted in A00066 and J99202; tests to distinguish increased destruction versus decreased production were not done. Bone density did not drop between week 6 and 10 in IL-7-treated
animals (Figure 8). This argues against
IL-7 causing osteoporosis. Moreover, bone biopsy was taken from J99202
before and at the end of IL-7 treatment and stained for
osteoclasts.45 No increase in osteoclasts was observed.
Lymphoma had not developed by 5 to 6 months after transplantation. In the 3 animals treated with IL-7 from the main experiment, this was documented by the absence of a mass on autopsies of M99149 and K99307 at 6 months and a CT scan of M99267 at 5 months after transplantation. Autopsies of J99202 and A00066 (5 and 6 months after transplantation) were also negative for a mass. Loss of fat or increased density of fatty tissue on CT was noted in 3 of 3 IL-7- and 0 of 3 placebo-treated animals (Figure 9). This was improved on the scans at
week 14. It was not associated with weight loss or clinical problems.
This could conceivably be due to T-cell infiltration of fatty tissue
because in one week 10 skin biopsy from an IL-7-treated animal that
happened to be deep enough to contain subcutaneous fat, the fat was
markedly infiltrated with T cells.
Two additional problems, possibly related to IL-7, were
noted
IL-7-neutralizing antibodies and IL-7 levels Neutralizing antibodies developed by week 10 in M99267, did not develop in M99149 (the best responder by CD4 counts filled circles in
Figure 1A), and were indeterminate in K99307 due to the presence of an
inhibitor of the growth of the IL-7-dependent cell line in all serum
samples, including those from before transplantation and from 6 weeks
after transplantation. Neutralizing antibodies also developed in J99202
and not in A00066.
Serum IL-7 levels increased approximately 1000-fold in the
IL-7-treated animals by the end of treatment, whereas the levels remained stable in the placebo-treated animals (Figure
11).
Here IL-7 was shown to increase CD4 T-cell counts in primates after hematopoietic cell transplantation. At the same time, potential side effects were noted, in particular, thrombocytopenia and a GVHD-like disease. The increase in CD4 T-cell counts occurred through the stimulation of peripheral expansion. Disappointingly, stimulation of de novo generation, which might broaden T-cell repertoire, was not observed. Nevertheless, stimulation of peripheral expansion alone might be of benefit. In mice, infections as well as tumors regressed as a result of IL-7-induced expansion of T cells.46-49 These results suggest that clinical trials are warranted, to test whether antiviral or antitumor activity of endogenous or infused T cells can be enhanced by IL-7. Our results suggest that patients participating in such trials should be carefully observed for hematologic toxicity and autoimmunity/GVHD. Patients with preexisting autoimmunity/GVHD should be excluded from such trials. The reason that IL-7 stimulated de novo generation in mouse models13,24,50 but not in the monkeys is unclear. The following possibilities exist: (1) A molecule other than IL-7 may be the homeostatic regulator of T-cell counts in primates. As T cells in the IL-7-treated baboons expanded, this putative regulator may have caused decreased T-cell generation de novo. (2) Neutralizing antibodies may have prevented the binding of IL-7 to IL-7 receptors on thymocytes. However, this is unlikely because there was no significant increase in TRECs in the animals that did not develop neutralizing antibodies (M99267 and A00066). (3) In bone marrow, IL-7 or the T cells infiltrating the marrow may have inhibited the production of thymocyte precursors or thymic dendritic cell precursors. (4) Filgrastim (G-CSF) given during the neutropenia associated with IL-7 treatment may have inhibited the generation of T cells.51 Interestingly, the animals that did not receive G-CSF during IL-7 treatment had higher increases in circulating TREC+ CD4 T cells (4.3-fold in K99307 and 5.2-fold in J99202) than the animals that during IL-7 treatment received G-CSF (1.4-fold, 2.3-fold, and no increase, respectively, in M99267, M99149, and A00066). Contrary to the original hypothesis, de novo generation may have been inhibited by IL-7, as suggested by lower TREC+ CD4 T-cell counts at 5 to 6 months after transplantation (Figure 1A) and lower thymic volume at 10 to 14 weeks after transplantation (Figure 2) in IL-7- versus placebo-treated animals. This is consistent with a recent observation that in normal macaques TREC+ T-cell counts declined during and after IL-7 treatment.52 B-cell counts were not increased by IL-7 in our monkeys. Whereas rodent
B lymphopoiesis is critically dependent on IL-7, our results as well as
those of other investigators suggest that primate B lymphopoiesis is
not significantly influenced by IL-7.52-55 In vitro production of B cells from marrow precursors requires IL-7 in
mice but not in humans.53 IL-7, IL-7R Toxicities, possibly attributable to IL-7, included a GVHD-like
disease and cytopenias. A GVHD-like disease occurs in about 20% of
patients receiving grafts with autologous CD34 cells.56 In
humans typically the disease develops within the first 2 months after
transplantation and is self-limited,56 whereas in our baboon it developed at 3.5 months after transplantation (within 1 week after starting IL-7) and was life-threatening. We have not
observed a GVHD-like disease developing at more than 2 months after
transplantation in more than 30 baboons receiving transplants of CD34
cells at our center over the last 10 years (without using IL-7). This
and the time of onset of the GVHD-like disease in the affected baboon
suggest that IL-7 may have caused the disease. Late cytopenias ( Twelve hours after the last dose of IL-7, IL-7 levels were around 10 ng/mL (3-4 logs above normal). Although we have not performed a formal pharmacokinetic study, this result suggests that in clinical trials IL-7 may need to be given only once a day or even less frequently, or that a lower dose may be effective. The optimal dose for human studies will need to be determined in phase 1 dose-escalation studies. IL-7 levels were very high (> 10 ng/mL 12 hours after the last dose of IL-7 treatment) even in the animals that developed IL-7-neutralizing antibody (M99267 and J99202). This suggests that the ELISA and the functional neutralization assay detected different IL-7 epitopes and that antigen-antibody complexes may have not been effectively cleared in animals treated with IL-7. In conclusion, several important differences in IL-7 effects between mice and primates have been identified. Similar to mice, IL-7 appears to hasten CD4 T-cell reconstitution in primates. Whereas both de novo generation and peripheral expansion are stimulated in mice, only the latter appears to be stimulated in primates. B lymphopoiesis is stimulated in mice but not primates. Autoimmune disorders were observed in our baboons but not in IL-7-treated mice (they were observed in IL-7 transgenic mice). Loss of fat or increased x-ray density of fat was observed in our baboons but not in mice. Osteoporosis developed in mice but not in baboons. Hematologic toxicity was significant in baboons but not in mice. These findings have important implications for the design of trials testing IL-7 in humans.
This work could not be done without the dedication of the staff of the University of Washington Regional Primate Research Center, in particular, Leslie Falch, Bruce Brown, Ed Novak, Mac Durning, Peggy Smith, Dr Judy Johnson, Dr David Anderson, Dr Steven Kelley, and Dr Maggie Gillen. The dedication and help of the computer tomogram staff was also essential, namely, Olivia Hicks, Mario Ramos, and Pat Manion. We also thank Dr Susan M. Ott for bone histology. We also thank Dr Michel Morre of Cytheris (Vanves, France) for providing the IL-7.
Submitted September 3, 2002; accepted January 6, 2003.
Prepublished online as Blood First Edition Paper, January 23, 2003; DOI 10.1182/blood-2002-08-2671.
Supported by National Institutes of Health grants HL69710, AI46108, CA18221, NCRR00166, DK56465, and DK47754, and a Fred Hutchinson Cancer Research Center (FHCRC) institutional pilot grant (funded by Bristol-Meyers-Squibb, New York, NY).
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
Reprints: Jan Storek, FHCRC, D1-100, 1100 Fairview Ave N, Seattle, WA 98109; e-mail: jstorek{at}fhcrc.org.
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Top
Abstract
Introduction
8 to day 
-mercaptoethanol.
Subsequently, the cells were starved of IL-7 for 24 hours (cultured in
the same media but without IL-7). The starved cells were transferred to a 96-well plate (100 000 cells/well) and cultured at 37°C in a humidified 5% CO2 atmosphere for 44 hours in the same
media, except for containing various concentrations of IL-7 and 15%
baboon serum instead of fetal bovine serum. A neutralizing
mouse-antihuman IL-7 monoclonal antibody (R & D Systems clone 7417.111)
was added to some wells at 2.5 µg/mL (positive control for
inhibition). After 44 hours the cultured cells were exposed to
3H-thymidine (1 µCi/well; 0.037 MBq/well) for 4 hours. The cells were lysed with water and DNA was harvested on a
filter (Packard-96 GF/C; Meriden, CT) using a semiautomated harvester
(Micromate 196; Packard). Counts per minute (CPMs) were determined in a
0.2 × 109/L) developed after
week 6 in 2 of 3 IL-7-treated and 0 of 3 placebo-treated animals.
Severe neutropenia did not develop in J99202 or A00066 animals.
-glutamyltransferase
12 times above baseline, alanine aminotransferase 2 times above
baseline, and bilirubin 3 times above baseline. The thymus was small,
barely visible, at autopsy and microscopically contained few
CD3+ cells (both CD4+ and CD8+);
the corticomedullary junction was indistinct. Spleen and mesenteric LNs
were moderately enlarged, whereas axillary and inguinal LNs were of
normal size. No germinal centers were present; most cells in the spleen
(both in the white and red pulp) and in the LNs were T cells (both
CD4+ and CD8+); B cells (CD20+)
were markedly decreased. Viral inclusions or syncytia were not seen in
any of the organs sampled. Original magnifications, × 30 (A) and × 50 (B-C).
2
months after transplantation) of presumed or documented autoimmune
etiology have been reported after human autografting.