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Stem Cells 2003;21:304-314 www.StemCells.com
© 2003 AlphaMed Press

In Utero Hematopoietic Stem Cell Transplantation in Nonhuman Primates: The Role of T Cells

Laurence E. Shieldsa,b, Lakshmi K. Gaurb,c, Mike Goughb, Jennifer Potterd, Aimee Sieverkroppd, Robert G. Andrewsb,d,e

a Department of Obstetrics and Gynecology, Division of Perinatal Medicine, University of Washington, Seattle, Washington, USA;
b National Primate Research Center, University of Washington, Seattle, Washington, USA;
c Puget Sound Blood Center, Washington Regional Primate Research Center, University of Washington, Seattle, Washington, USA;
d Clinical Research Division, Fred Hutchinson Cancer Center;
e Department of Pediatrics, University of Washington, Seattle, Washington, USA

Key Words. Nonhuman primate • Fetal transplantation • Fetal therapy

Laurence E. Shields, M.D., Department of Obstetrics and Gynecology, Division of Perinatal Medicine, Box 356460, University of Washington, Seattle, Washington 98105-6460, USA. Telephone: 206-543-3714; Fax: 206-616-9479; e-mail: lshields{at}u.washington.edu


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
In utero transplantation of hematopoietic stem cells is a promising treatment for immune and hematologic diseases of fetuses and newborns. Unfortunately, there are limited data from nonhuman primates and humans describing optimal transplantation conditions. The purpose of this investigation was to determine the effect of T-cell number on engraftment and the level of chimerism after in utero transplantation in nonhuman primates. CD34+ allogeneic adult bone marrow cells, obtained from the sire after G-CSF and stem cell factor administration, were transplanted into female fetal recipients. The average CD34+ cell dose was 3.0 x 109/kg (range, 9.9 x 108 to 4.4 x 109) and the T-cell dose ranged from 2.6 x 105 to 1.1 x 108/kg. Chimerism was determined in peripheral blood subsets (CD2, CD13, and CD20) and in progenitor cell populations by using polymerase chain reaction. Chimerism was noted in seven of eight live-born animals. The level of chimerism in the progenitor population was related to the fetal T-cell dose (r = 0.64, p < 0.02). At the lowest T-cell dose (2.6 x 105/kg), no chimerism was detected. As the T-cell dose increased to 106–7/kg, the level of chimerism increased. Adjusting the T-cell dose to 1.1 x 108/kg resulted in fatal graft-versus-host disease (GVHD). The results of this study emphasize the importance of T cells in facilitating donor cell engraftment and in producing GVHD in fetal nonhuman primates. Some animals achieved levels of chimerism in the marrow hematopoietic progenitor cell population that would likely have clinical relevance. However, the levels of chimerism in peripheral blood were too low for therapeutic benefit. Further studies are needed to test methods that are likely to enhance donor cell engraftment and peripheral blood levels of donor cells.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Although most inheritable diseases affecting hematopoietic stem cells and lymphohematopoietic progeny can be cured by allogeneic bone marrow transplantation, in some individuals, irreversible organ damage has occurred prenatally or early in life, making postnatal therapy less efficacious. In utero transplantation of allogeneic hematopoietic stem cells has been proposed as an alternative therapy that may potentially reduce the morbidity and mortality associated with postnatal bone marrow transplantation, as well as potentially reducing the deleterious effects that occur prior to birth.

Data from naturally occurring [1–4] and experimental animal models [5–9] of in utero hematopoietic stem cell transplantation suggest that therapeutic trials in the human fetus should be successful. Unfortunately, to date, the only successful attempts of in utero hematopoietic stem cell transplantation in humans have been in fetuses with severe immunologic defects [10–14]. The reasons for the lack of success in human fetal stem cell transplantation have not been elucidated. The composition of the donor cell graft, the number of cells transplanted, and the ages of the fetal recipients have only been systematically studied in the ovine (sheep) model. While many animal models provide valuable insights toward our understanding of the biology and success of cell-based therapies, they frequently do not completely mimic the biologic and clinical situation present in the human. Nonhuman primates show similar ontogeny of immune and hematopoietic systems and have been a relevant animal model for the study of postnatal bone marrow transplantation, providing data that have been directly transferred to the human.

Therefore, the purpose of this investigation was to determine the effect of T-cell number on engraftment and the level of chimerism after in utero transplantation of purified, haploidentical CD34+ allogeneic adult bone marrow cells in a nonhuman primate model.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animal Selection
The use of nonhuman primates in this study was an approved research protocol through the University of Washington Animal Care Committee and meets AAALAC guidelines. Pregnant baboons (Papio species, n = 2 [F96-118, M98-179]) and macaques (Macasa nemestrina, n = 9) were identified through time-mating or pregnancy surveillance programs using ultrasound at the Washington Regional Primate Research Center. Initial studies were done in baboons, but due to the greater availability of pregnant macaques, the majority of animals used were M. nemestrina. After pregnant animals were identified, the gestational age was determined by ultrasound [15]. Fetal weights of normal fetuses autopsied at various gestational ages were estimated based on previously published data from our center [16]. From these data, a fetal growth chart was established and used to calculate estimated fetal weights prior to each injection. All transplanted fetuses were female. Fetal sex was determined from amniotic fluid cells obtained by ultrasound-guided amniocentesis using polymerase chain reaction (PCR) for a Y-chromosome-specific DNA sequence (see below). The absence of a PCR signal for Y-specific DNA with a positive control PCR (actin) was used to determine that the fetus was female.

Fetal Transplantation
Donor cells were injected into the fetal abdomen under ultrasound guidance using a 25-g spinal needle (Becton Dickinson; Franklin Lakes, NJ; http://www.bd.com) at 0.34–0.38 gestation. This gestational age was chosen based on studies showing an optimal transplant time of ~0.38–0.46 gestation in fetal sheep [8] and others showing no chimerism in nonhuman primates when fetuses were transplanted at >0.44 gestation (baboons) [17] and >0.42 gestation (cynomolgus monkeys) [18]. The success of each fetal injection was confirmed by observing a small echogenic focus at the injection site representing a small amount of air purposely injected with the donor cells and by the presence of a small amount of abdominal fluid (ascites) [19]. Each fetus received two (animals F96-118, M98-179, K97-177, M00-202) or three (animals M99-080, M00-311, M00-233, M01-035, K00-025, M00-074, L00-053) injections of donor cells at weekly intervals. The initial injection was performed at 0.36 ± 0.01 gestation. The donor cells were fresh for the first and third injections, and in all cases, the second injection was from a cryopreserved aliquot.

Donor Cell Preparation and Monoclonal Antibodies
All donor cells were from the sire of the treated fetus allowing the use of PCR for Y-chromosome-specific DNA to determine chimerism (see below). In all but one case, the donor was treated with G-CSF (100 µg/kg, Amgen, Inc.; Thousand Oaks, CA; http://www.amgen.com) and stem cell factor (SCF, 50 µg/kg, both factors kindly provided by Dr. Graham Molineux, Amgen, Inc.] for 4–5 days prior to bone marrow harvest. Bone marrow was aspirated from proximal humeri and distal femurs into heparinized syringes. Marrow buffy coat cells were isolated and CD34+ cells were enriched using the mouse IgM anti-CD34 monoclonal antibody 12.8 (25 µg/ml), anti-mouse IgM monoclonal antibody-conjugated magnetic beads, and VS+ magnetic columns according to the manufacturer’s instructions (Miltenyi Biotec; Auburn, CA; http://www.miltenyibiotec.com), as previously described [20]. Aliquots of enriched cells were labeled with either phycoerythrin (PE)-conjugated goat anti-mouse IgM (Immunotech; Marseille, France) or mouse monoclonal anti-CD2 (S5.2, BD Pharmingen; San Diego CA; http://www.bdbiosciences.com/pharmingen) conjugated with fluorescein isothiocyanate (FITC) to enumerate CD34+ and CD2+ cells using flow cytometric analysis (Cell Quest software; Becton-Dickinson; Mountain View, CA; http://www.bd.com). For purposes of cell sorting and analysis, a cell was considered to be positively labeled if it had more fluorescent intensity greater than 99% of the cells stained with the appropriate isotype control monoclonal antibody (H12C12), IgM antimouse Thy 1.2 [21], and IgG (G1) (Becton Dickinson). Post-CD34 enrichment, the donor cell population was 76% ± 12% (mean ± standard deviation [SD]) CD34+ and 0.51% ± 0.45% CD2+.

T-cell depletion was performed on CD34-enriched cells using the magnetic bead system (Dynal; Oslo, Norway; http://www.dynal.no). One animal received a donor cell graft that was subjected to T-cell depletion. Briefly, the CD34+-enriched cells were washed twice and suspended at 1 x 108 cells/ml in phosphate buffered solution-human AB serum (PBS-HABS; Gemini Bio-Products; Woodland, CA; http://www.gembio.com). The cells were then incubated with a mouse IgG anti-CD2 monoclonal antibody (S5.2; BD Pharmingen) at 4°C for 20 minutes. The cell suspension was then washed twice in PBS-HABS and incubated with anti-mouse IgG-coated magnetic beads (Dynal) at a concentration of 5 beads per cell. Cells and beads were then mixed and pelleted at 1,200 rpm for 5 minutes and incubated at 4°C for 5 minutes. The pellet was gently resuspended, and cells adsorbed to magnetic particles were removed by applying a magnet to the side of the tube. Nonadsorbed cells were collected in the supernatant, washed twice with PBS/HABS, and then separated again using a magnet to remove residual magnetic particles. The frequency of CD2+ cells in the nonadsorbed population was determined by flow cytometry. T-cell depleted CD34+-enriched cells were used to engraft one fetus (M99-080).

T-cell enrichment was performed on peripheral blood obtained from the donor prior to initiation of growth factor stimulation. Enriched CD2+ cells were then given to two fetuses (to increase the total number of donor T cells in the donor cell graft). Briefly, after red cell lysis, the cells were incubated with a mouse IgG anti-CD2 monoclonal antibody (S5.2, BD Pharmingen) at 4°C for 20 minutes. The cell suspension was then washed twice in PBS-HABS, incubated with anti-mouse IgG-conjugated magnetic beads, and enriched using VS+ magnetic columns according to the manufacturer’s instructions (Miltenyi Biotec), as previously described [20]. After enrichment, the cells were 90% CD2+. The cells were then aliquoted and frozen in 90% fetal calf serum (FCS) and 10% dimethylsulfoxide. At each donor cell injection, the same number of T cells was added back to the preparation. Two animals (L00-53 and M00-074) received T-cell-enriched grafts. L00-053 died in utero at 102 days, and tissue maceration precluded detailed study of chimerism or tissue histology. M00-074 died near term with histologic evidence of graft-versus-host disease (GVHD).

Peripheral Blood Cell Processing from Transplanted Animals
Peripheral blood cells from transplanted animals were isolated as buffy coat preparations, as previously described [22]. The cells were labeled with FITC- or PE-conjugated monoclonal antibodies (CD2, CD4, CD13, CD20) or appropriate isotype controls (Becton Dickinson). Cells within the appropriate light scatter gate were then sorted using a Vantage fluorescence-activated cell sorter (FACS, Becton Dickinson) into purified CD2, CD4, CD13, and CD20 cells. The purity of these cells was >99%. The cells were placed into aliquots of 1,000–5,000 cells for DNA extraction.

Progenitor Cell Assays
Double-layer agar colony-forming cell (CFC) assays were used to evaluate chimerism in the hematopoietic progenitor population, as previously described [23, 24]. Cells cultured in double-layer colony assays were stimulated with multiple hematopoietic growth factors (interleukin-3 [IL-3], IL-6, SCF, GM-CSF, and G-CSF, each at 100 ng/ml [kindly provided by Dr. Graham Molineux, Amgen, Inc.]) and erythropoietin (Epo) at 4 IU/ml (Amgen). Fetal cord blood and bone marrow cells were plated at 10,000 cells/plate. The presence of BFU-E and CFCs granulocyte-macrophage was determined by colony morphology after 12–14 days of incubation at 37°C in an atmospheric mixture of 5% O2, 5% CO2, and 80% nitrogen. Colonies were picked using a dissecting microscope.

To further reduce any risk of contamination from non-CFCs within the soft agar culture system, we also grew CFCs from CD34+ marrow cells that were deposited as single cells by FACS directly into wells of 96-well "U" bottom plates (Corning Inc.; Corning, NY; http://www.corning.com). These individual cells were cultured in 170 µl of {alpha} minimal essential medium (GIBCO BRL; Grand Island, NY; http://www.invitrogen.com) supplemented with 25% fetal bovine serum, 1% bovine serum albumin (fraction V; Sigma; St. Louis, MO; http://www.sigmaaldrich.com), 50 µg/ml gentamicin sulfate (Gemini Bio-Products Inc.; Calabasas, CA; http://www.gembio.com), and multiple hematopoietic growth factors (IL-3, IL-6, SCF, G-CSF, GM-CSF, and Epo) at the same concentrations used in agar-based cultures. Single-cell cultures were incubated at 37°C in an atmospheric mixture of 5% O2, 5% CO2, and 80% nitrogen for 10–14 days. The wells were screened using a dissecting scope, and colonies were collected for DNA extraction

Chimerism: PCR for Male-Specific Sequence
To isolate genomic DNA, cells from individual colonies plucked from agar or microtiter wells and FAC-sorted cells (CD2, CD4, CD13, and CD20) were mixed with 10 µl PBS, 90 µl nanopure water, and 2 µl (200 µl/ml) of protease K (Roche Diagnostic Corp.; Indianapolis, IN; http://www.roche-diagnostics.com) and heated using a thermal cycler (MJ Research; Watertown MA), as previously described [25]. DNA samples that were not immediately used were stored at -80°C. PCR was performed using primers previously described by Reitsma et al. [26], RhM3 (5' GAA AGA ACA TAA AGG ACC TA 3') and RhM4 (5' GGT AGA ATT AAT ATG ACC 3') (Fred Hutchinson Cancer Research Center Biotech Service Center) to amplify a 174-b, Y-chromosome-specific DNA sequence. This sequence could be amplified from DNA of male macaques and baboons but not from female macaques, baboons, or male or female humans. Controls, assayed as part of each PCR run, included DNA isolated from 100,000 cells that were 100% female; 100% male; and mixtures of female:male cells of 10:1, 100:1, 1,000:1, and 10,000:1, as well as water. In our lab, this methodology required between 10–100 male cells to be present to obtain a positive PCR signal and would detect as few as 0.1%–0.01% male cells mixed with cells from a normal female of the same species (baboons or macaques). As a control for DNA integrity, PCR was performed on DNA from the same sample for an actin sequence (approximately 300 b) using primers (actin 1 = 5'TCC TGT GGC ATC CGC GAA ACT3' and actin 2 = 5' GAA GCA TTT GCG GAC GAT 3') [25].

Mixed Lymphocyte Cultures
Mixed lymphocyte cultures (MLCs) were performed from peripheral blood mononuclear cells. Triplicate samples of 1 x 105 responder cells and 1 x 105 irradiated (3,000 rad) stimulator cells were cultured in RPMI medium (Invitrogen; Grand Island, NY) and 10% FCS, in 96-well round-bottom microplates. DNA synthesis was measured on day 6 of culture after a 12-hour pulse with 3H thymidine. The proliferative response was quantitated by 3H thymidine incorporation (c.p.m.). Values from each cultured set are presented as the percent relative response (RR). Relative response = ([experimental MLC – autologous MLC]/[unrelated MLC – autologous MLC] x 100) [27, 28].

Graft-Versus-Host Disease
Clinical GVHD was defined as runting in combination with other clinical findings (rhinorrhea, loss of fur, diarrhea, or persistent skin lesions). Suspected in utero GVHD was made on the basis of hydrops fetalis and confirmed postnatally by histologic exam [18]. Pathologic GVHD was defined as lymphocytic infiltrates noted in skin, intestine, liver, or other tissues.

Statistical Analysis
The level of chimerism (the proportion of male colonies) was determined using data from PCR on DNA of both individually picked and pooled colonies. By using the equation p =1-(1-r/n)1/m (where p = probability of any clone being positive, r = number of PCR positive pools, n = number of pools tested, and m = number of colonies per pool), the frequency of PCR-positive colonies could be estimated [29]. The variance of p is defined by the equation [1-(1-p)m]/[nm2(1-p)m-2]. Because such a sensitive assay could be susceptible to contamination leading to false-positive reactions, control experiments used known mixtures of male and female marrow cells. For those experiments, marrow cells from normal male and female animals were mixed at ratios (male:female) of 25:75, 10:90, 1:99, and 0.1:99.9. The cells were then plated at 30,000 cells/35-mm dish. After 14 days of culture, individual colonies were picked and collected in pools of 1, 2, 5, or 15 colonies per pool. The DNA was extracted and subjected to PCR (Table 1Go). The rate of observed chimerism did not differ significantly from that expected from ratios of mixed cells.


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  		Table 1. PCR estimates of the frequencies of male progenitors in CFCs grown from known mixtures of male and female marrow
 

   RESULTS
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 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Transplanted Animals
Eleven animals were transplanted at a mean of 0.36 ± 0.01 gestation. Seven animals were live born. Two animals were born dead, but tissue was usable for data analysis. Thus, nine animals were available for analysis. The other two animals died in utero. One of the fetuses was noted to be dead at the time of the third donor cell injection (77 days), presumably as a result of injection trauma, and one was delivered as a fetal demise after premature rupture of the fetal membranes at 0.60 gestation (102 days). The nine animals that were available for analysis received an average of 3.0 x 109 CD34+ cells (range, 9.9 x 108 to 4.4 x 109 CD34+ cells/kg). The total T-cell dose ranged from 2.6 x 105 to 1.1 x 108/kg. The one animal that did not demonstrate chimerism in cord blood or bone marrow obtained shortly after birth had received the lowest number of donor T cells (2.6 x 105/kg). The animal that received the highest T-cell dose (1.1 x 108/kg), where donor T cells were added back to the cell preparation, delivered as an intrauterine demise at term with severe hydrops fetalis. Based on the condition of the fetus at the time of delivery (lack of maceration or skin sloughing), the animal appeared to have died shortly before birth. Pathological examination of that animal showed hydropic changes and lymphocytic infiltrates in skin and subcutaneous tissue consistent with GVHD [18]. Attempts to grow cells from the fetal liver, bone marrow, and cord blood were not successful, and we were not able to quantitate the level of chimerism. The six animals that are still alive show no evidence of abnormal growth or clinical evidence of GVHD.

Chimerism in Transplanted Animals
Chimerism was detected in cord blood cells or bone marrow cells and peripheral blood cells from seven of the eight animals (87%) in the first month of life (Table 2Go). To further evaluate chimerism, we studied hematopoietic progenitors from cord blood and bone marrow grown either in agar or as single CD34+ cells in liquid medium and analyzed by PCR (Fig. 1Go). The seven chimeric animals had donor cells detected in their hematopoietic progenitor populations (Table 3Go). There was a strong relationship between the total number of T cells given to the fetus and chimerism within progenitor populations (r = 0.64, p < 0.02; Fig. 2Go). Chimerism was not detected in bone marrow or cord blood of the animal that received the lowest T-cell dose (105/kg). As the number of T cells in the donor cell graft was increased from 106 to 107 cells/kg, the level of chimerism also increased. The T cell dose was intentionally adjusted to approximately 108 CD2+ T cells in two animals. The first received 1.1 x 108/kg CD2+ cells along with 2.5 x 109/kg CD34+ cells. The dam of this fetus had premature rupture of the fetal membranes at 103 days gestation. The animal was delivered after the fetus was noted to be dead. Histologic evaluation of the fetus was difficult due to the macerated condition of the fetus as a result of intrauterine infection, and we could not exclude GVHD. Chimerism studies were uninformative (no PCR result from control actin). The second animal transplanted with 1.1 x 108 CD2+ T cells developed severe hydrops fetalis, and histologic evaluation demonstrated lymphocytic infiltrates in the skin and subcutaneous tissue consistent with severe GVHD.


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  		Table 2. Engraftment: presence of male (donor) cells in cord blood, peripheral blood, and bone marrow within the first month of life
 


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  		Figure 1. PCR gel from animal M00-311. The top lanes are controls (female, male, and aliquots of 100,000 cells in dilutions of 1:10, 1:100 and 1:1,000 of male:female macaques cells). W = water. The lower lanes show positive male control, water, and 1 x 105 bone marrow cells aliquoted from a single marrow sample (lanes 1–7). The brightness of the PCR signal is consistent with values between 1:100 and 1:1,000 (male:female dilutions).

 

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  		Table 3. Chimerism of hematopoietic progenitor cells
 


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  		Figure 2. A) Data showing the relationship between the initial level of chimerism and the total number of CD2+ T cells/estimated fetal weight in the donor cell graft. B) Regression analysis showing the relationship between the total number of T cells/kg given to the fetus and the level of chimerism in the progenitor cell population (r = 0.64; p < 0.02).

 
We evaluated six animals to determine whether donor cells contributed progeny to lymphoid and myeloid lineages in the peripheral blood. Peripheral blood CD2, CD4, CD13, and CD20 cells were isolated by FACS. In four animals, donor-derived cells were detected in aliquots of purified T and B lymphocytes, as well as myeloid cells (Table 4Go). In a fifth animal, donor cells were detected in myeloid and B-lymphoid blood cells. A sixth animal with <1% progenitor chimerism had donor cells detected only in the B-lymphoid lineage. While the life span of lymphocytes is relatively long, and lymphocytic cells could simply be cells that were contaminants of our original donor cell preparation, the short life expectancy of the mature myeloid cells in the circulation suggests that donor hematopoietic cells matured into these progeny. Assuming a need for 10–100 male cells to achieve a positive PCR, one can estimate the frequency of donor cells circulating in the peripheral blood of our chimeric animals. This estimate varied from 0.1-0.002. Based on the fact that we did not observe a positive PCR result in all aliquots tested from any animal or in any lineage, one would have to assume that the level of peripheral blood chimerism was at the lower end of this estimate. Thus, despite our ability to achieve relatively high levels of donor cells in the hematopoietic progenitor cell population, the frequency of circulating donor cells was relatively rare.


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  		Table 4. Chimerism within multiple peripheral blood lineages
 
The stability of the donor graft in the hematopoietic progenitor population in marrow (CFCs and individual CD34+ cells cultured from marrow) was studied in six animals (Table 3Go). Follow-up varied from 2–24 months. In four of the animals, the levels of progenitor cell chimerism increased. In one animal, the level of chimerism was increased at 13 months of age but, by 24 months of age, was barely detectable (0.01%), and in another animal, the level of chimerism in bone marrow decreased from 6% at birth to 1.3% at 8 months of age. The temporal changes in progenitor chimerism did not appear to correlate with the number of CD34+ or CD2+ cells transplanted.

Donor-Specific Tolerance
Mixed lymphocyte culture was used to determine whether tolerance was induced after in utero transplantation in six of the animals. In four of the six animals tested, the recipient response to the donor was markedly less than that of the dam and unrelated animals (Table 5Go). In two of the animals, the MLC responses of the neonate to the sire and dam were equivalent. One of these animals did not have evidence of donor chimerism and the other animal initially demonstrated chimerism in the progenitor population (2.7%), but when MLC testing was carried out at 2 years of age, the level of chimerism was low (0.1%) and the response to the sire (donor) was similar to that of the dams.


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  		Table 5. Recipient MLC relative responses to donors, dams, and unrelated animals by mixed lymphocyte testing
 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Successful postnatal engraftment is correlated with the total CD34+ cell dose and requires providing sufficient donor T cells. Although the number of animals in this study was relatively small, it supports a donor T-cell number (CD2+ cells) on the order of 106-107/kg estimated fetal weight. In one animal, where the total T-cell dose was 108/kg, the animal died with massive hydrops fetalis with lymphocytic infiltrates on histologic exam, consistent with GVHD. Similar physical findings in the fetus have been previously reported when the marrow cell preparation was felt not to have been sufficiently T-cell depleted [18].

The only other systematic evaluation of the effect of donor T-cell number on the success of in utero transplantation has been in the fetal sheep model. Although the fetal sheep model has been criticized for not providing data that translate to successful human fetal transplantation, the findings from studies of fetal sheep are remarkably similar to those of nonhuman primates [30]. In the sheep model, a T-cell dose of 106 to 107/kg resulted in engraftment in the majority of animals without GVHD. At a T-cell dose of 108/kg, the majority of the animals engrafted, but the rate of GVHD was unacceptably high (75%–100%). In the human fetus, intrauterine death also occurred from "overwhelming engraftment" [31] when the T-cell dose exceeded 108/kg. While the fetus has been assumed to be "preimmune" [32, 33], it is probably more correct to term the fetus "early immune," as immunocompetent cells are present very early in the fetus, suggesting that it is not truly devoid of immune function [34–41]. If one assumes that the fetus has some immune function, it is not surprising that a relatively large number of donor T cells are needed to ensure successful engraftment, presumably through a donor-versus-host effect. Future studies utilizing cellular conditioning or modulating the fetal immune system at the time of in utero transplant should be undertaken to see if fetal immune function is inhibiting engraftment of allogeneic cells after in utero hematopoietic stem cell transplantation.

Four other investigators have published studies of in utero hematopoietic stem cell transplantation in nonhuman primates. Harrison et al. [42] published a series of five in utero transplants in rhesus monkeys (M. mulatta) using donor cells from randomly selected allogeneic fetal livers. Durable engraftment was noted in the hematopoietic progenitor population of the four animals tested. Direct comparison of this study with our results is difficult due to the different cell source and the lack of specific information related to the number of T cells in the graft. Evaluation of cell source in the allogeneic sheep model has suggested that the highest level of engraftment occurs after transplantation with fetal-liver-derived cells [8]. Comparing levels of chimerism achieved in our study with those of Harrison et al. suggests that if there is any potential advantage to fetal liver as a donor cell source, this might be overcome by the use of larger numbers of CD34+ cells and T cells obtained from growth-factor-stimulated adult bone marrow.

More recently, Cowan et al. [28] published a larger series of in utero transplants in rhesus monkeys using haploidentical, T-cell depleted adult marrow as donor cells. The animals were transplanted at an average of 44 days (0.25 gestation) or 61 days (0.37 gestation). In both gestational age groups, the level of progenitor chimerism was generally low (<1%). However, two animals demonstrating long-term engraftment showed higher levels of chimerism. Karyotype analysis at approximately 3–6 months of age demonstrated 2/100 and 2/50 metaphase samples positive for donor cells. Our study differed in both the cell type (growth-factor-stimulated and CD34+-enriched versus T-cell-depleted bone marrow) and total CD34 cell dose. We also used multiple time points to inject our donor cells based on data from fetal sheep and neonatal mice, suggesting that giving the same number of donor cells through multiple injections increases the level of chimerism [8, 43]. It is unclear which of these factors resulted in the different outcomes between these two studies. Our total T-cell dose and that calculated from Cowan et al. were similar and varied from 105-107/kg estimated fetal weight. The use of very large numbers of growth-factor-stimulated CD34+ cells may have increased the probability of successful in utero therapy by a variety of mechanisms. G-CSF-stimulated donor cells have also been shown to have different adhesion molecule expression (particularly ß1 integrin and L-selectin expression), cytokine production patterns [44], and more T cells with type-2 cytokine production patterns [45]. In postnatal transplantation, the total CD34 cell dose has been correlated with engraftment kinetics [46, 47] and graft failure, in the case of transplants with umbilical cord blood. It is not surprising that in the absence of pretransplant conditioning and in the fetal setting, where the frequency and proliferative capacity of endogenous fetal CD34+ cells is very high [24, 48], a very large number of donor cells will be needed for transplantation.

In addition to achieving donor cell engraftment in the hematopoietic progenitor population, we also achieved multilineage engraftment with donor cells noted in peripheral blood T cells, B cells, and CD13 myeloid cells. The presence of donor cells in the short-lived CD13+ myeloid cell population suggests that those cells were not simply present in the original cell population. Despite the limitations of our present donor cell detection system, it is clear that, even with relatively high levels of donor progenitor cells in the marrow, the number of circulating mature progeny from donor hematopoietic cells was infrequent. Similar findings of lower peripheral blood relative to marrow chimerism have been noted in the sheep xenogeneic, and to a lesser degree, the allogeneic fetal transplantation model [32, 49]. The reasons for the higher levels of chimerism in the progenitor population than those noted in peripheral blood are not completely clear but may be related to incomplete matching between stromal and stem cell major histocompatibility complex (MHC), as well as differences in fetal stromal and adult stem cell interactions. In vitro culturing of mouse hematopoietic stem cells with MHC-matched and -mismatched stromal cells has shown enhanced hematopoiesis and secretion of cytokine growth factors from matched stromal cells relative to mismatched stromal cells [50]. In addition, in the fetal sheep model of hematopoietic stem cell transplantation, the use of adult stromal cells with adult-derived stem cells significantly increased the level of donor cell progeny in the peripheral blood [32, 49]. Alternatively, the low levels of peripheral blood chimerism may be related to relative quiescence of nonhuman primate hematopoietic stem cells [51]. These factors should be taken into consideration in future studies of allogeneic fetal transplantation.

The stability of the donor cell graft was evaluated by progenitor cell assays in six animals. Chimerism was noted to increase in four animals, to increase and then decrease to nearly undetectable levels in one animal, and to decrease over time in one animal. The direction of change was not related to the number of CD34+ or CD2+ cells given to the fetuses. It is possible that these changes merely reflect changes in the number of donor cells cycling in the marrow at any one time. Other factors that mediated the direction of change have not yet been elucidated. If in utero therapy is going to be useful in the clinical setting, the graft must persist and achieve levels of chimerism that are curative for the disease being treated. One method that could potentially be used to improve the level of chimerism in the setting of declining donor cell chimerism or low levels of chimerism is postnatal donor cell booster dosing. In both mice [6, 52] and sheep [53] models, postnatal booster dosing has been associated with an improvement in the level of chimerism. One animal, whose progenitor chimerism declined from 6% to 1.3% by 8 months of life and had MLC results suggesting donor-specific tolerance, was boosted with donor marrow cells from its original donor. This animal was given a single infusion of 3.8 x 108 T-cell-depleted marrow cells harvested after mobilization of the donor with G-CSF and SCF. Three months after infusion, 18/146 (12.3%) individually cultured CD34+ marrow cells were of donor origin (unpublished preliminary data). The results in this one animal, while consistent with reports of postnatal booster dosing in neonatal mice and sheep, will require confirmation in additional animals. It will be important to evaluate the immune consequences of in utero hematopoietic stem cell therapy and postnatal boosting.

Our data suggest that relatively high numbers of donor T cells are needed to ensure engraftment in fetal nonhuman primates. The precise number of donor cells needed in the graft is not clear but appears to be in the range of 1 x 106–7/kg estimated fetal weight. We were also able to demonstrate a relatively high percentage of donor cells in the CD34+ progenitor population. However, peripheral blood chimerism was very low and certainly not at a level that would be expected to treat most diseases that should be amendable to in utero therapy. Future studies will need to address methods of enhancing donor cell engraftment and, more specifically, donor cell proliferation.


   ACKNOWLEDGMENT
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
We appreciate the kind donation of recombinant human G-CSF and SCF from Dr. Graham Molineux from Amgen Corporation. This work was supported by National Institutes of Health grants HL62422-02 and RR00166.


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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Received October 22, 2002; accepted for publication February 1, 2003.



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