www.elsevier.com/locate/yclim

Clinical Immunology

Interleukin-7 improves reconstitution of antiviral CD4 T cells$

Hailing Lu, Zhao Zhao, Tomas Kalina, Thurman Gillespy III, Denny Liggitt, Robert G. Andrews,

David G. Maloney, Hans-Peter Kiem, Jan Storek1

Fred Hutchinson Cancer Research Center and University of Washington, Seattle, WA, United States

Received 11 June 2004; accepted with revision 10 August 2004

Abstract

We evaluated whether long-term (2 months) administration of interleukin-7 (IL7) hastens immune recovery in baboons rendered severely

lymphopenic by total body irradiation and antithymocyte globulin (ATG). Four baboons were treated with recombinant baboon IL7 and three

baboons with placebo. Median CD4 T cell count at the end of IL7/placebo treatment was higher in the IL7-treated animals (2262 vs. 618/Al,P = 0.03). This appeared to be a result of peripheral expansion rather than de novo generation. Median cytomegalovirus (CMV)-specific

IFNg-producing CD4 T cell count at the end of IL7/placebo treatment was higher in the IL7-treated animals (122 vs. 1/Al, P = 0.03). All

animals were pretransplant cytomegalovirus-seropositive. One animal died at the end of IL7 treatment; necropsy showed extensive T cell

infiltration of kidneys and lungs. In conclusion, IL7 stimulates the expansion of CD4 T cells, including functional antiviral cells. Clinical

risk–benefit ratio needs to be evaluated.

D 2004 Elsevier Inc. All rights reserved.

Keywords: T cell reconstitution; Interleukin-7; Hematopoietic cell transplantation; Antiviral immunity; Cytomegalovirus; T lymphocytes; Immunodeficiency

diseases; Viral diseases; Cytokines

Introduction

CD4 T-lymphopenia in hematopoietic cell transplant

recipients and HIV-infected patients is associated with

infections [1–3], including cytomegalovirus (CMV) infec-

tions [4,5]. IL7 has been shown to increase CD4 T cell

counts in both murine and nonhuman primate models [6–

11]. Both de novo generation of T cells (from hematopoietic

cells, typically in the thymus) and peripheral expansion of

existing T cells were reported to be stimulated by IL7 in

mice [9]. However, only peripheral expansion was stimu-

lated in the two primate studies reported, that is, Fry et al.’s

[10] study using normal and SIV-infected macaques and our

study using irradiated baboons [11]. It is possible that de

1521-6616/$ - see front matter D 2004 Elsevier Inc. All rights reserved.

doi:10.1016/j.clim.2004.08.008

$ Supported by National Institutes of Health Grants No. HL68710 and

CA18221.1 Present address: University of Calgary, Health Science Centre, 3330

Hospital Drive NW, Calgary, Alberta, Canada T2N 4N1. Fax: +1 403 283

1267.

E-mail address: jstorek@fhcrc.org (J. Storek).

novo generation was not observed in the two primate studies

because the treatment was too short (10 days in Fry et al.’s

study and 28 days in our study), because xenogeneic IL7

was used or because only moderately lymphopenic monkeys

were treated.

The present study was thus performed for the following

reasons: (1) to study the effect of IL7 in severely

lymphopenic monkeys, as stimulation of de novo generation

by IL7 was not observed in T cell-replete mice [12]; (2) to

administer baboon instead of human IL7, as baboon–

antihuman IL7 antibodies [11] or, less likely, relative

incompatibility between human IL7 and baboon IL7

receptor could diminish the IL7 effect; (3) to treat the

animals for a longer period (8 weeks) than in the previous

studies; as thymocyte precursor to T cell differentiation

takes at least 12 days [13], only prolonged treatment might

result in a detectable rise of recent thymic emigrants,

represented by T cell receptor excision circle-containing

(TREC+) T cells; (4) to add data on potential side effects of

IL7 in primates, as clinical trials of IL7 in patients with

iatrogenic or HIV-induced lymphopenia are being planned;

114 (2005) 30–41

H. Lu et al. / Clinical Immunology 114 (2005) 30–41 31

(5) most important, to determine whether the IL7-stimulated

increase in the total CD4 T cell counts might be functionally

relevant. To evaluate this, we treated CMV-seropositive

baboons and asked whether higher anti-CMV CD4 T cell

counts were achieved in the IL7 than placebo-treated

animals. Reconstitution of anti-CMV CD4 T cells is

essential for the reconstitution of anti-CMV CD8 T cells

(reviewed by Gratama and Cornelissen [14]), and increased

numbers of CMV-specific CD4 T cells have been associated

with decreased incidence of symptomatic CMV disease

[15].

Materials and methods

Study design and animals

Seven male baboons (Papio cynocephalus anubis), age 2

years (equivalent to 5–7 human years [16–18]), underwent

total body irradiation (TBI, 10.2 Gy) followed by autolo-

gous transplantation of CD34 cells marked with yellow

fluorescent protein (YFP) gene. Before starting IL7/placebo

treatment, T cells were further depleted by antithymocyte

globulin (ATG, thymoglobulin, 5 mg/kg intravenously on

days 32, 34, 36, and 39 posttransplant, kindly donated by

Sangstat, Fremont, CA). Importantly, in vivo ATG causes

apoptosis of circulating splenic and lymphonodal T cells but

does not affect thymocytes, probably because of limited

access of ATG to the thymus [19]. Three baboons received

placebo, and four baboons received recombinant baboon

IL7 (kindly donated by Cytheris, Vanves, France) 80 Ag/kgonce a day subcutaneously from days 42 to 98 after

transplantation. In the Results, day 42 is referred to as

bimmediately pre-IL7/placebo treatmentQ and day 98 as

bweek 8 of treatment.Q Lyophilized recombinant baboon IL7

was reconstituted with a diluent containing 4 mg/ml sucrose

and 125 mM sodium citrate, pH 6.2. The same diluent was

used as placebo. Pretransplant, the animals were sero-

positive for CMV, seronegative for Babesia and STLV-I, and

tuberculin test-negative. They were housed and cared for at

the University of Washington Regional Primate Research

Center under conditions approved by the American Asso-

ciation for the Accreditation of Laboratory Animal Care.

The study was approved by the UW Animal Care and Use

Committee and the FHCRC Institutional Review Board.

Transplantation and posttransplant treatment

Filgrastim (G-CSF), 100 Ag/kg daily, and stem cell factor

(SCF), 50 Ag/kg daily, were given subcutaneously from days

8 to 4 to increase the number of CD34 cells in the marrow.

On day 3, bone marrow was harvested into preservative-free

heparin from both humeri and femora. CD34 cells were

immunomagnetically enriched to approximately 80% purity,

using antibody 12.8 and goat antimouse IgM-coated

microbeads (Miltenyi, Auburn, CA). Transduction of

CD34 cells with an oncoretrovirus carrying the enhanced

YFP gene was done as described [20]. Approximately 50%

transduced and 50% untransduced CD34 cells were infused.

(The utility of transplanting the YFP gene-transduced CD34

cells was limited, as sustained engraftment of the transduced

cells was achieved in only four of the seven animals.) After

total body irradiation (10.2 Gy in two fractions at 0.07 Gy/

min), the cells were reinfused on day 0. Supportive care

included irradiated whole blood transfusions for thrombo-

cytopenia and prophylactic antibiotics (acyclovir and fluco-

nazole from days 0 to 112, ceftazidime + gentamycin +

vancomycin from days 0 to 41 and levofloxacin from days

41 to 112 posttransplant).

Enumeration of blood mononuclear cell subsets

Mononuclear cells (MNCs) were separated from EDTA-

anticoagulated blood using density gradient (Ficoll, density

1.073 kg/l) centrifugation. The cells, suspended in flow

buffer (PBS with 1% bovine serum albumin and 0.1%

sodium azide), were stained with fluorochrome-conjugated

monoclonal antibodies and analyzed by five-color flow

cytometry. The following antibodies were used: CD3 from

Biosource (clone FN18), CD4 from BD Biosciences (clone

SK3), CD8 from Beckman-Coulter (clone B9.11), CD11a

from Beckman-Coulter (clone 25.3.1), CD14 from Beck-

man-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 + 1% potassium bicarbonate + 0.4% EDTA). No

fixation was done. Cells were resuspended in flow buffer

and kept on ice until flow cytometry, which occurred within

12 h from blood drawing. FACSCalibur (BD Biosciences)

flow cytometer was used for data acquisition. List-mode

data were analyzed using WinList software (Verity Software

House, Topsham, ME). The 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. 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 (LN) were obtained immediately before

starting IL7 (right axilla) at week 8 of treatment (left axilla)

and approximately 3 months later (right axilla). Axillary

LNs were chosen as they were distant from the IL7/placebo

injection sites (thighs). The LNs were teased in RPMI with

10% fetal calf serum. The resulting cell suspension was

H. Lu et al. / Clinical Immunology 114 (2005) 30–4132

filtered through a 70-Am nylon cell strainer. Percentages of

cell subsets were determined by flow cytometry as in the

blood.

Thymic biopsy was done in one placebo-treated and two

IL7-treated animals at week 8 of treatment, using upper

sternal split approach under general anesthesia. Approx-

imately 3000 mm3 of thymic tissue was removed and

processed for flow cytometry as for lymph nodes and for

histology as described below.

CMV serology

The presence of antibodies against baboon CMV was

tested by Simian Diagnostic Laboratory (San Antonio, TX)

using rapid dot-immunobinding assay [21]. In brief, virus or

control antigens were spotted on the nitrocellulose sheet and

allowed to absorb for 30 min at room temperature. Then the

nitrocellulose sheet was submerged in 5% nonfat milk in

PBS to block nonspecific binding for 1 h followed by

washing with PBS-Tween (0.1%). After test sera were

applied over the test antigen dots, the nitrocellulose sheet

was incubated for 1 h at 378C and then washed three times

in PBS-Tween. Then staphylococcal protein A-conjugated

horseradish peroxidase was used to detect primate IgG.

Following incubation and washing, the substrate, 4-chloro-

1-naphthol-hydrogen peroxide, was used, and a blue spot

was developed as a positive result. Controls consisted of

uninfected cell culture antigens and known positive and

negative sera.

Intracellular cytokine staining for detection of CMV-specific

CD4 T cells

We used CMV lysate to stimulate baboon MNCs. CMV

lysate was prepared using MRC5 human lung fibroblast cell

line (ATCC CCL-171) infected with baboon CMV, strain

PGH-01 (ATCC VR-1530) or OCOM4-37 (kindly provided

by Dr. Earl Blewett, Oklahoma State University). After

infection had progressed to N90% cytopathic effect, the cells

were harvested and subjected to multiple freeze–thaw

cycles. Then the cells were washed twice in glycine-

buffered saline, and the pellets were sonicated on ice. The

resulting suspension of cell organelles/particles was centri-

fuged at 2000 rpm for 10 min, and the supernatant was

harvested. The supernatants from both CMV strains were

mixed and heat-inactivated at 568C for 30 min. Protein

concentration was determined with a BCA protein assay kit

(Pierce, Rockford, IL). Aliquots of the CMV lysate were

stored in �808C. Control lysate with the same protein

concentration was prepared from uninfected MRC5 cells.

One million PBMC in 1 ml of RPMI 1640 supplemented

with 2 mM glutamine, 25 mM HEPES, 10% fetal bovine

serum (Hyclone, Logan, UT) were incubated with CMV

lysate (5 Ag protein/ml), uninfected MRC5 lysate (5 Agprotein/ml, negative control), or staphylococcal enterotoxin

B (1 Ag/ml, Sigma, positive control) in the presence of anti-

CD28 and anti-CD49d antibodies (0.5 Ag/ml each, BD

Biosciences, San Jose, CA) overnight at 378C in a

humidified atmosphere. Brefeldin A (10 Ag/ml; Sigma-

Aldrich, St. Louis, MO) was added 2 h after the initiation of

the culture. After the overnight incubation, cells were

treated with 1 mM EDTA for 10 min at RT. Then the cells

were sequentially fixed and permeabilized using FACS

lysing solution and FACS permeabilizing solution (BD

Biosciences). The cells were then stained with anti-IFNg-

phycoerythrin (clone 45.15, Beckman Coulter [Immuno-

tech], Miami, FL) or anti-TNFa-phycoerythrin (clone

Mab11, BD Biosciences), anti-CD3-PerCPCy5.5 (clone

SK34-2, BD Biosciences), anti-CD8 APCCy7 (clone SK1,

BD Biosciences), and anti-CD4-APC (clone SK3, BD

Biosciences) for 30 min at room temperature in the dark.

After final wash with flow buffer, data acquisition was done

on LSR II cytometer (BD Biosciences). At least 200,000

events were acquired. Data were analyzed using FlowJo

software (Treestar, San Carlos, CA).

Detection of CMV by PCR

DNA was extracted from 200 Al serum using QIAamp

DNA Blood Mini Kit (Qiagen, Valencia, CA). The

following primers were used: 5V-TACGTCATTGG-

TACCCTCC-3V(forward) and 5V-GGAGTACTGCCAATG-TACTA-3V(reverse) [22]. Each reaction contained 2.5 U

platinum-Taq polymerase (Invitrogen, Carlsbad, CA), 2.5

mM MgCl2, 500 uM dNTPs, 500 nM each primer, 20 Alextracted DNA (of total 100 Al yielded by the QIAamp

kit) in a total volume of 50 Al. The conditions of each

reaction were 1 cycle of 958C for 9 min, followed by 40

cycles of 948C/60 s; 588C/60 s; 728C/60 s, and finally 1

cycle of 728C for 9 min. PCR products were separated

on 2% agarose gel and visualized by ethidium bromide

staining. As a positive control, we used DNA extracted

from the CMV lysate (derived from baboon CMV-

infected MRC5 cells). As negative control, we used

DNA extracted from baboon lymphocryptovirus (Epstein-

Barr virus-like virus)-transformed baboon B lymphoblas-

toid cells.

T cell receptor excision circle (TREC) assay

FACS-sorted CD4 T cell pellets of 200,000 cells were

lysed in 40 Al of lysis buffer containing 100 Ag/ml

proteinase K and 10 mM Tris–HCl (pH 8.0). The 5V-nuclease (Taqman) assay for the baboon ay signal joint

TREC [11] was performed on 5 Al of cell lysate using

forward primer, 5VCACATCCCTTTCAACCATGCT3V,reverse primer, 5VGCCAGCTGCAGGGTTTAGG3V, and

probe FAM-ACGCATTTGGTTTTTGTAAAGGTGCT-

CACT-QSY7 (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 958C for

H. Lu et al. / Clinical Immunology 114 (2005) 30–41 33

5 min, then 958C/30 s, and 608C/1 min 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 in 5 Al of cell lysate, which corresponds to

25,000 cells) was calculated using the ABI7700 software.

The absolute count of TREC+ CD4 (CD8) T cells (per

microliter) was calculated as the TREC level (per 25,000

cells) multiplied by the absolute CD4 T cell count (per

microliter) and divided by 25,000.

Computed tomography (CT)

Volumes of the thymus, the spleen, and the lymph

nodes and the density of the cancellous bone of L3, L4,

and L5 vertebrae were measured as described [11]. The

calculated density of the torso phantom ranged from 94.6

to 96.7 mg/cm3, suggesting adequate reproducibility. The

volume of the kidneys was determined analogously to the

thymus. Briefly, the area of kidneys (mm2) was determined

on each slice, using IV contrast images and GE Pathspeed

software (GE Medical Systems). The area was multiplied

by the slice thickness (1.25 mm) to get the volume at that

slice level (mm3). Kidney volume (of both kidneys

together) was calculated as the sum of the volumes at

each slice level. Increase of volume of an organ was

calculated as the volume at the end (week 8) of IL7/

placebo treatment minus the volume immediately pre-IL7/

placebo treatment.

Histology

Using standard techniques, 10% neutral buffered for-

malin-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) or CD20 (clone L26, Dako). OTC-

embedded snap-frozen tissues were immunostained with

CD4 (clone 1F6, Novocastra, Newcastle upon Tyne, UK) or

CD8 (clone G10.1, kind gift of Dr. Bing Hu, University of

Washington). Slides were analyzed by one pathologist

(D.L.) blinded to IL7 versus placebo treatment. Bone

marrow cellularity was estimated from the H and E-stained

sections 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.

IL7 levels and IL7 neutralizing antibodies

Serum concentration of IL7 was determined by ELISA

using the kit for human IL7 (R and D Systems, Minneapolis,

MN). In case of values exceeding the concentration of the

highest standard, diluted serum was used.

IL7 neutralizing antibodies were determined by the

inhibition of the growth of IL7-dependent cells (PB1) as

described [11].

Statistics

To compare the counts of CD4 T cells or their subsets at

the end of IL7/placebo treatment for the animals that

survived until the week 8 evaluation, we used the average

count from weeks 7 and 8 of the IL7/placebo treatment. For

the animal that died 1 day before the week 8 evaluation

(Z00158), we used the count from week 7. Significance of

differences was tested by the Mann-Whitney–Wilcoxon

rank sum test. As the hypothesis was that the cell counts

were higher in the IL7-treated animals, one-tailed P value

was used.

Results

IL7 treatment improved the reconstitution of CD4 T cells but

not CD8 T cells

After total body irradiation and further T cell depletion

by ATG, the baboons were severely lymphopenic.

Immediately before starting IL7/placebo treatment, median

CD4 T cell count in the blood was 3/Al, and lymph nodes

were severely lymphocyte-depleted (composed of mostly

histiocytes). At the end of IL7/placebo treatment (weeks

7–8), median CD4 T cell count in the blood was higher in

the IL7-treated animals (2262 vs. 618/Al, P = 0.03) (Fig.

1). The median number of weeks it took to reach

individual pretransplant CD4 T cell count was 2.5 weeks

for IL7-treated versus 8 weeks for placebo-treated animals

(P = 0.03). The improved CD4 T cell reconstitution

appeared to be primarily, if not entirely, due to the

stimulation of peripheral expansion, as the count of

TREC+ CD4 T cells (per microliter of blood which

reflects thymic output irrespective of peripheral expansion

[not the percent of TREC+ cells among CD4 T cells which

is influenced by peripheral expansion]) was not signifi-

cantly higher in IL7 than placebo-treated animals (Fig. 1).

Both CD45RAhigh and CD45RAlow/� CD4 T cell counts

were significantly higher at the end of treatment (weeks

7–8) in IL7 than placebo-treated animals (P = 0.03) (Fig.

1). This, together with the fact that TREC+ CD4 T cell

counts were not significantly higher in the IL7-treated

animals, suggests that IL7 induced expansion of both

naive as well as memory/effector cells. The naive cells

expanded by IL7 likely included cells generated de novo

since the time of transplant; as in the four animals with

sustained YFP marking (two IL7- and two placebo-

treated), there was a trend toward higher YFP+ CD4 T

cell counts at the end of treatment (weeks 7–8) in the IL7-

than placebo-treated animals (median 294 vs. 68/Al, datanot shown).

Fig. 1. Counts per microliter blood (top and center) or percentages (bottom) of CD4 T cells or their subsets in IL7-treated (black thin lines) and placebo-treated

(gray thick lines) animals. On each x-axis, bPre-TxQ denotes pretransplant, bPre-IL7Q denotes immediately pre-IL7/placebo treatment, and subsequent numbers

denote weeks after starting IL7/placebo treatment. The period of IL7 treatment is indicated in each graph by the black horizontal bar. The TREC+CD4Tcell counts

could not be reliably enumerated immediately before and in the first 3 weeks of IL7/placebo treatment due to low total CD4 counts; b0.01Q is arbitrarily displayedfor the immediate pretreatment time point. BMdenotes bonemarrow, and LN denotes lymph nodes. In both organs, the percentages are among total nucleated cells

and were determined by flow cytometry. For a clearer interpretation of this figure, the reader is referred to the figure in the Web version of this article.

H. Lu et al. / Clinical Immunology 114 (2005) 30–4134

The increased blood CD4 T cell counts in IL7-treated

animals were not due to redistribution of CD4 T cells from

tissues to blood; as in the bone marrow of IL7-versus

placebo-treated animals, there was a trend toward a higher

percentage of CD4 T cells (Fig. 1) and a higher overall

marrow cellularity (median 62.5% vs. 55% nonfat cells

among fat plus nonfat cells at week 4 and 72.5% vs. 40% at

week 8). In the lymph nodes at week 8, the percentage of

CD4 T cells was similar (Fig. 1), while there was a trend

toward larger lymph node volume in IL7- than placebo-

treated animals (Fig. 2). These results suggest that IL7

treatment increased the number of total body CD4 T cells.

The conclusion that IL7 stimulated peripheral expansion

rather than de novo generation was further supported by the

determination of the volumes of thymi, spleens, and lymph

nodes (Fig. 2). There was a trend toward higher median

increase of splenic volume (30,063 vs. 13,914 mm3) and

lymph node volume (1150 vs. 714 mm3) but not thymic

volume (191 vs. 2682 mm3) from the start to the end of IL7

versus placebo treatment. This was consistent with the

Fig. 2. Volumes of thymus, spleen, and lymph nodes (LN) in IL7-treated

(black) and placebo-treated (gray) animals pretransplant (Pre-Tx),

immediately pre-IL7/placebo treatment (Pre-IL7), and 8 and 20 weeks

after starting IL7/placebo. Each LN volume represents the sum of

volumes of one right axillary LN, one left axillary lymph node, one

pelvic LN, one right inguinal LN, and one left inguinal lymph node. IL7

treatment period is indicated by the horizontal black bar. The animals

subjected to thymic biopsy at week 8 (after the CT scan) had the

following symbols: black asterisks, black circles, and gray circles. Note

that, at the last follow-up, five of six animals reached a higher thymic

volume than pretransplant; the one animal that did not reach pretransplant

thymic volume (gray circles) may have not done so due at least in part to

the biopsy at week 8. For a clearer interpretation of this figure, the reader

is referred to the figure in the Web version of this article.

H. Lu et al. / Clinical Immunology 114 (2005) 30–41 35

necropsy findings on the animal that died 1 day before the

week 8 evaluation (Z00158); its spleen and lymph nodes

were larger, and its thymus was smaller than normal (per

pathologist blinded to the treatment—D.H.L.). Histologi-

cally (by H and E stain) and flow cytometrically (by

percentages of CD4�CD8�, CD4+CD8+, CD4+CD8�, and

CD4�CD8+ thymocytes), the thymus of Z00158 as well as

the thymi of the other two IL7-treated and one placebo-

treated animals subjected to thymic biopsy at week 8

appeared normal. The conclusion that IL7 stimulated

peripheral expansion rather than de novo generation was

also supported by the fact that there appeared to be no

difference in the percentage of TREC-containing cells

among lymph node CD4 T cells at week 8. This was

determined in two IL7- and two placebo-treated animals,

and the average was 11% versus 12%, respectively.

IL7 had no significant effect on blood counts or lymph

node percentages of CD8 T cells, B cells, monocytes, or NK

cells (data not shown).

IL7 treatment increased the number of CMV-specific CD4 T

cells

Symptomatic herpesviral disease occurs in CD4

lymphopenic patients, particularly those with low counts

of herpesvirus-specific CD4 T cells [3,15,23]. We asked

whether herpesvirus-specific CD4 T cells are increased

with IL7. CMV was used as a prototype herpesvirus

because baboon CMV infection of baboons closely

resembles human CMV infection of humans [24] and

because we could identify baboon CMV-specific CD4 T

cells in baboons as CD4 T cells producing IFNg when

stimulated with baboon CMV antigens (Fig. 3, top).

Because total CD4 T cell counts were low, CMV-specific

CD4 T cells could not be reliably enumerated immedi-

ately pre-IL7/placebo treatment (zero values are arbitrarily

displayed in Fig. 3, bottom left) and could not be always

enumerated in the first 6 weeks or treatment, especially

in the placebo-treated animals. Nevertheless, a complete

set of data was available for the end of treatment (weeks

7–8). At that time point, median CMV-specific CD4 T

cell counts were significantly higher in IL7 than placebo-

treated animals (median 122 vs. 1/Al, P = 0.03). Thus,

IL7 treatment increased anti-CMV CD4 T cells.

The IFNg-producing CD4 T cells typically produced

also TNFa (Fig. 3, bottom right). In 18 specimens

studied (from pretransplant and various time points

posttransplant), percent CD4 T cells producing TNFa

(alone or together with IFNg) correlated with the percent

of CD4 T cells producing IFNg (alone or together with

TNFa) (Spearman rank correlation coefficient R = 0.90,

P b 0.001). A similar correlation was found in macaque

CMV-infected macaques [25]. Therefore, it is likely that

counts of not only IFNg-producing anti-CMV CD4 T

cells but also TNFa-producing anti-CMV CD4 T cells

were higher in IL7 than placebo-treated animals.

Fig. 3. IL7 treatment increased CMV-specific CD4 T cells. At the top, examples of CD4 versus IFNg dotplots from the end of treatment (week 7 or 8) are

displayed, showing the trend toward higher percentages of CMV-specific IFNg-producing CD4 T cells (among total CD4 T cells) in IL7- than placebo-

treated animals. The bNormalQ blood was obtained pretransplant. Blood MNCs were stimulated with CMV lysate prepared from infected MRC5 cells and

processed as described in the Materials and methods. Stimulation with uninfected MRC5 lysate resulted in less than 0.05% IFNg-producing CD4 cells (not

shown); this was subtracted when calculating the absolute number of CMV-specific CD4 T cells displayed below. In each dotplot, the gate is set on CD4

lymphocytes, and 10,000 events are displayed. At the bottom left, the absolute numbers of CMV-specific CD4 T cells in IL7 (black)- and placebo (gray)-treated

baboons are shown. IL7 treatment period is indicated by the horizontal black bar. CMV-specific cells could not be reliably enumerated immediately pre-IL7/

placebo treatment; b0Q is arbitrarily displayed for each animal. At the bottom right, a typical TNFa versus IFNg dotplot gated on CD4 lymphocytes is

displayed, showing that most IFNg CD4 T cells produced also TNFa. For a clearer interpretation of this figure, the reader is referred to the figure in the Web

version of this article.

H. Lu et al. / Clinical Immunology 114 (2005) 30–4136

After transplantation in humans, recovery of CMV-

specific T cells is boosted by CMV reactivation (detection

of CMV in posttransplant blood using a technique that

usually does not detect CMV in blood from normal

seropositive individuals, for example, pp65 immunostain-

ing) [26–28]. In our experiment, CMV reactivation could

have occurred in the IL7-treated and not in the placebo-

treated baboons. If true, the higher counts of CMV-specific

CD4 T cells in the IL7-treated baboons could be due to

CMV reactivation and not necessarily due to IL7. Contrary

to that, reactivation of CMV during IL7/placebo treatment

was documented in all (both IL7- and placebo-treated)

baboons (Fig. 4). Therefore, the increased number of CMV-

specific CD4 T cells in the IL7-treated animals was likely

due to IL7.

Potential toxicity of IL7

One animal (Z00158) died unexpectedly at the end of

IL7 administration (1 day before the scheduled week 8

Fig. 5. Extensive lung (top) and kidney (center) infiltration with CD3+ cells

(brown). This was found on the necropsy of animal Z00158 that died 1 day

before week 8 evaluation. In the lungs, the T cells were localized primarily

around blood vessels (asterisks), but also around bronchioles and in the

parenchyma. In the kidneys, the T cells were localized also primarily

around blood vessels (asterisks), but also around tubules and glomeruli.

Among the surviving animals, kidney enlargement (bilateral) at week 8 of

treatment was found in the IL7-treated but not the placebo-treated animals

(bottom). The kidney volume is a composite volume of right and left

kidneys. Image (top and center) information: Nikon E-400 microscope,

Nikon 10� lens with 0.25 numerical aperture, digital imaging medium,

Nikon Coolpix 5000 camera with Nikon software, Microsoft Photo Editor

(for adjustment of general brightness and contrast only—no other

operations performed).

Fig. 4. Reactivation of CMVoccurred in all seven baboons (four IL7-treated

baboons, upper gels; and three placebo-treated baboons, lower gels). Viral

DNA was detected in serum by PCR immediately before IL7/placebo

treatment (bPre IL7Q) and at most time points during IL7/placebo treatment

(weeks 1–7) but not pretransplant (bPre TxQ). CMV lysate (derived from

baboon CMV-infected MRC5 cells) was used as the positive control (+) and

baboon Epstein-Barr virus-transformed baboon B lymphoblastoid cells

were used as the negative control (�). N/A indicates serum not available.

H. Lu et al. / Clinical Immunology 114 (2005) 30–41 37

evaluation). The only clinical symptom or sign noted was

anorexia in the last week of life, attributed to mild renal

insufficiency (serum creatinine rose from baseline 0.5–1.0

mg/dl to maximum 1.6 mg/dl). On necropsy, macroscopic

findings included large lymph nodes and spleen, small

thymus, marginally enlarged kidneys with pale, radial

streaks in the cortex, and firm and nonfloating lungs.

Microscopically, most organs were infiltrated with CD3+

cells. The infiltration was most prominent in the lungs and

the kidneys (Fig. 5, top). Approximately one half of the cells

infiltrating the kidneys and the lungs were CD4+ and one

half CD8+. No inclusions or syncytia were noted, suggesting

autoimmune rather than viral etiology. Of the total of nine

baboons we have treated with IL7, death associated with T

cell infiltration of multiple organs occurred in two animals

(A00066 reported in Ref. [11] and Z00158 reported here).

This complication may have resulted from overtreatment, as

we continued to give IL7 to animals per protocol even after

they have reached a high CD4 T cell count. The death

occurred in two of five animals that reached a CD4 T cell

count of greater than 2,500/Al and none of four animals that

did not reach CD4 T cell count of 2500/Al. CD8 T cell

counts and the deaths appeared to be unrelated.

Because of the renal dysfunction of Z00158 premortem

with the postmortem finding of T cell infiltration of the

kidneys reported here and because of a mild kidney

enlargement from immediately pre-IL7 to the end of 4-

week treatment noted on CT in all five animals treated

with IL7 in our previous experiment [11] (unpublished),

we retrospectively measured kidney volumes of the three

surviving IL7-treated and the three placebo-treated animals

in the present experiment (Fig. 5, bottom). Median

increase in the kidney volume was 31,155 mm3 in the

IL7-treated group compared to �4301 mm3 in the

Fig. 6. Hematocrit and platelet counts in IL7 (black)- and placebo (gray)-

treated baboons. The high hematocrit immediately pre-IL7/placebo treat-

ment is a result of overtransfusion (using whole blood for thrombocyto-

penia) in the first 6 weeks after transplant. The sudden increases in

hematocrit in the animal denoted by black triangles between 2 and 3 weeks

and in the animal denoted by gray squares between 3 and 4 weeks into IL7/

placebo treatment resulted from blood transfusions. The sudden drop in

hematocrit in the animal denoted by gray squares between weeks 14 and 15

resulted from babesiosis, and the subsequent rise in hematocrit coincided

with doxycycline treatment. As in Fig. 1, bPre-TxQ denotes pretransplant,

bPre-IL7Q denotes immediately pre-IL7/placebo treatment, and subsequent

numbers denote weeks after starting IL7/placebo treatment. The period of

IL7 treatment is indicated by the black horizontal bar. For a clearer

interpretation of this figure, the reader is referred to the figure in the Web

version of this article.

Fig. 7. Density of cancellous bone in IL7 (black)- and placebo (gray)-

treated baboons. Average density of the cancellous bone of vertebral bodies

L3, L4, and L5 showed no decrease during IL7 treatment. IL7 treatment

period is indicated by the horizontal black bar. For a clearer interpretation of

this figure, the reader is referred to the figure in the Web version of this

article.

H. Lu et al. / Clinical Immunology 114 (2005) 30–4138

placebo-treated group. Concurrent rise in serum creatinine

was not observed. The kidney volume returned close to

baseline within 3 months after discontinuation of IL7. We

speculate that the kidney enlargement was due to T cell

infiltration, although histology was only available for

Z00158.

Thrombocytopenia and possibly neutropenia and anemia

were associated with IL7 treatment in the previous experi-

ment. In this experiment, we observed a trend toward lower

hematocrit and platelet counts (Fig. 6) but not neutrophil

counts (data not shown) in IL7- than placebo-treated

animals. Perhaps, the hematologic toxicity in this experi-

ment was not as obvious as in the previous experiment

because one of the placebo-treated animals had an unusually

poor engraftment (gray squares in Fig. 6).

Osteoporosis due to stimulation of osteoclasts developed

after only 20 days of treatment with IL7 in mice. [29]. In our

previous baboon experiment, treatment with IL7 for 28 days

was not associated with decreased bone density. In the

present experiment, treatment for 56 days was also not

associated with decreased bone density (Fig. 7).

Lymphoma did not develop in the three IL7-treated

animals that did not die unexpectedly. This was determined

by both CTand necropsy at 3 months after discontinuation of

IL7.

IL7 levels and IL7 neutralizing antibodies

Median serum IL7 level (of all seven animals) rose from

6 pg/ml pretransplant to 27 pg/ml immediately pre-IL7/

placebo treatment, reflecting endogenous response to the

severe CD4 lymphopenia (Fig. 8). In the IL7-treated

animals, IL7 levels approximately 3 logs higher than at

baseline (pretransplant) were reached within 1 week and

were maintained throughout the remaining 7 weeks of

treatment. In the placebo-treated animals, IL7 levels

returned to baseline within approximately 1 month.

IL7 neutralizing antibodies may have theoretically been

generated in the IL7-treated animals, although the IL7 was

species-specific, as human cytokine neutralizing antibodies

have developed in humans treated with erythropoietin,

GM-CSF and thrombopoietin [30–32], and IL7 might

stimulate antibody production by stimulating helper T

cells. Therefore, sera from all four IL7-treated baboons

Fig. 8. Serum levels of IL7 in IL7 (black)- and placebo (gray)-treated

baboons. As in Fig. 1, bPre-TxQ denotes pretransplant, bPre-IL7Q denotesimmediately pre-IL7/placebo treatment, and subsequent numbers on the x-

axis denote weeks after starting IL7/placebo treatment. The period of IL7

treatment is indicated by the black horizontal bar. For a clearer

interpretation of this figure, the reader is referred to the figure in the Web

version of this article.

H. Lu et al. / Clinical Immunology 114 (2005) 30–41 39

collected at the end of IL7 treatment were assayed for the

presence of IL7 neutralizing antibodies. None were

detected.

Discussion

This study shows that, even in the setting of severe

lymphopenia and with 8-week treatment (longer than

previously reported), IL7 does not substantially stimulate

thymopoiesis. The positive finding of this study is that

IL7 administered to primates expands anti-CMV CD4 T

cells that are functional (produce IFNg and TNFa when

stimulated). This gives hope that IL7 administered to

patients will expand antiviral or antitumor CD4 T cells

capable of producing IFNg and TNFa. IFNg- or TNFa-

producing CD4 T cells can directly kill virus-infected or

tumor cells [15,33–44]. Virus/tumor-specific CD4 T cells

may also facilitate protection against viral disease or

cancer indirectly by sustaining virus/tumor-specific cyto-

toxic CD8 T cells (reviewed in Ref. [14]). Major

application of IL7 will likely be in immunotherapy. For

example, after high-dose chemo/radiotherapy for cancer

with autologous transplantation of CD34 cells (purified to

minimize the likelihood of reinfusing tumor cells), T cells

specific for the cancer and for viruses that may cause

disease after autologous CD34 cell transplantation like

CMV [45] or Epstein-Barr virus [46] could be infused

and expanded in vivo by the administration of IL7. In

herpes simplex virus-infected mice treated with antiviral

T cells, IL7 treatment resulted in 20-fold reduction in

viral load compared to controls treated with antiviral T

cells alone [47].

Regarding potential toxicities of IL7, it is reassuring that

even after 8-week treatment, lymphoma (detectable as a

mass on CT or necropsy) or osteoporosis did not develop,

and anemia and thrombocytopenia appeared to be mild.

However, the two deaths associated with T cell infiltration

of multiple organs, occurring in nine IL7-treated baboons,

are of concern. In both cases, the organs were infiltrated

with both CD4 and CD8 T cells, suggesting against

monoclonal/malignant lymphoproliferation. Viral etiology

is also unlikely as no inclusions or syncytia were noted on H

and E-stained sections of the most involved organs

(intestines in the first case and lungs and kidneys in the

second case) or other organs. Direct autoimmune damage of

organs by the infiltrating T cells is a plausible mechanism; in

the first case, the T cells were concentrated in the areas of

damaged intestinal crypts (Fig. 10 in Ref. [11]). Alter-

natively, the IL7-expanded T cells may have secreted other

cytokines, leading to a bcytokine storm.Q This would be

reminiscent of interleukin-2 toxicity which appears to be

mediated through the release of secondary cytokines like

TNF, IFNg, interleukin-1, and interleukin-6 [48]. The latter

mechanism is less likely as in normal monkeys treated with

IL7 (up to 1200 Ag/kg/day for less than 1 month) for the

determination of IL7 toxicity, no increase in serum levels of

TNFa, IFNg, or interleukin-6 was observed (M. Morre,

personal communication, March 2004). Despite the fact that

the pathogenesis of the death associated with T lympho-

proliferation has not been elucidated, the important obser-

vation is that it may be related to the degree of the IL7-

induced CD4 T cell expansion. The deaths occurred in two

of five animals whose CD4 T cell count during IL7

treatment exceeded 2500/Al and in none of four animals

with lower maximum CD4 T cell count during IL7

treatment. Among 21 normal 2-year-old baboons in which

we determined CD4 T cell counts, the two highest counts

were 2005 and 2130/Al. Our suggestion for the design of

clinical trials is to monitor CD4 T cell counts during IL7

treatment and either discontinue IL7 or proceed with

extreme caution after the CD4 T cell count has exceeded

the upper normal limit.

In conclusion, given the potential for increasing the

number of functional antiviral or antitumor T cells,

clinical testing of IL7 is indicated. Analogous to the

fact that life-threatening side effects of erythropoietin

(e.g., occlusion of a cerebral vessel) occur in patients

reaching high erythrocyte counts and those of filgrastim

(e.g., dyspnea associated with pulmonary X-ray infil-

trates) occur in patients reaching high granulocyte counts,

it is possible that life-threatening side effects of IL7 may

occur in patients reaching high CD4 T cell counts.

H. Lu et al. / Clinical Immunology 114 (2005) 30–4140

Acknowledgments

We thank Monja D. Metcalf, Roxanne Velez, Julia

Morris, and Renee Kendall for excellent technical assis-

tance. This work could not be done without the dedication

of the staff of the University of Washington National

Primate Research Center, in particular, Michael Gough,

Leslie Falch, Ed Novak, Mac Durning, Peggy Smith, Judy

Johnson, Dr. David Anderson, Dr. Steven Kelley, and Dr.

Maggie Gillen. The dedication and help of the computer

tomogram staff were also essential, namely, Olivia Hicks,

Mario Ramos, and Pat Manion. We also greatly acknowl-

edge Drs. Michel Morre and Renaud Buffet of Cytheris for

providing us with the recombinant baboon IL7.

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