Graduate School of Pharmaceutical Sciences, University of Tokyo, Tokyo 113-0033, Japan
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ABSTRACT |
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Repeated administration of recombinant human erythropoietin (rhEPO) causes upregulation of receptor-mediated tissue uptake by the spleen (Kato, M., H. Kamiyama, A. Okazaki, K. Kumaki, Y. Kato, and Y. Sugiyama. J. Pharmacol. Exp. Ther. 283: 520-527, 1997). To discover whether such upregulation is due to an increase in target cells, the numbers of colony-forming unit erythroids (CFU-E) and burst-forming unit erythroids (BFU-E), the precursor of CFU-E, were measured in the spleen after rhEPO treatment. The uptake clearance of 125I-labeled rhEPO by the spleen was almost proportional to the number of CFU-E, suggesting that the upregulation is due to an increased number of CFU-E. When growth cells were metabolically labeled with [3H]thymidine in vivo, the radioactivity in bone marrow fell significantly after rhEPO treatment, whereas that in the spleen increased significantly. A cell-fractionation study using Percoll revealed that the radioactivity in the BFU-E fraction of splenic cells increased initially after rhEPO treatment, followed by an increase in radioactivity in the CFU-E fraction with a concomitant reduction in radioactivity in the BFU-E fraction. These results demonstrate that EPO stimulates the migration of BFU-E from bone marrow to spleen, followed by its differentiation into CFU-E in the spleen. In conclusion, the upregulation observed in the spleen is due to its stimulatory effect on the migration of BFU-E.
migration; receptor-mediated endocytosis; cytokine
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INTRODUCTION |
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RECENT INVESTIGATIONS have identified several types of hematopoietic growth factors, such as erythropoietin (EPO), granulocyte colony-stimulating factor (G-CSF), and granulocyte macrophage colony-stimulating factor, and attempts have been made to clarify their physiological roles. The recombinant products of these proteins are used to treat several hematological diseases. EPO is a 34-kDa glycoprotein that is mainly produced by the kidney, and it stimulates the proliferation and differentiation of colony-forming unit erythroids (CFU-E) (14). At present, recombinant human EPO (rhEPO) is used to treat anemia in patients with end-stage renal disease whose EPO production is low. The pharmacological target of EPO is CFU-E, which is the precursor of erythrocytes and on which EPO receptors are predominantly expressed.
To understand the physiological role of these biologically active proteins, it is important to investigate the mechanism of their distribution and elimination in the body. We have previously reported that bone marrow and spleen make a major contribution to the overall elimination of the G-CSF derivative nartograstim (15-17) and rhEPO (7) from blood circulation, whereas hepatotropic growth factors such as epidermal growth factor and hepatocyte growth factor are eliminated mainly by the liver (18, 23). Thus the target organs are the clearance organs for these types of proteins. This may be due to the major role played by receptor-mediated endocytosis (RME) in their distribution and elimination (16, 18, 23, 24, 27). The pharmacokinetics of these cytokines exhibits nonlinearity because of saturation of their receptors and/or downregulation of the receptor density on the target cell surface in the presence of excess ligand.
We have shown that repeated doses of rhEPO produce a change in the pharmacokinetics of rhEPO because of upregulation of EPO receptors. After rhEPO treatment, the hematocrit exhibited a significant correlation with the sum of the tissue uptake clearance (CLup) by bone marrow and spleen in both rats and mice (7, 8). These findings suggest that pharmacological receptors play a role in determining the magnitude of CLup. The CLup by the spleen is markedly increased by such rhEPO administration, whereas that by bone marrow is only slightly increased in both rats and mice (7, 8). Thus the upregulation of CLup is tissue specific. Repeated administration of nartograstim has also been reported to cause marked upregulation of G-CSF receptors. It has been suggested that this upregulation is due to an increase in the number of granulocyte-expressing G-CSF receptors in the spleen (16). On the other hand, the mechanism of tissue-specific upregulation of EPO receptors remains to be clarified. The present study was performed to answer the following questions. 1) Is the upregulation of EPO receptors in spleen due to an increase of the number of target cells (CFU-E)? 2) What is the mechanism for the difference in the degree of upregulation between bone marrow and spleen?
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MATERIALS AND METHODS |
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Materials. rhEPO was produced using Chinese hamster ovarian cells transfected with an expression vector harboring the human EPO cDNA at Production Technology Laboratories, Chugai Pharmaceutical (Tokyo, Japan). 111In-labeled indium chloride, 125I-labeled sodium iodide (17.4 Ci/mg), and [3H]thymidine were obtained from Amersham (Amersham, Buckinghamshire, UK). Diethylenetriamine-N,N,N',N'',N'''-pentaacetic acid anhydride (DTPA) was obtained from Dojindo (Kyoto, Japan). Iodo-Gen (1,3,6-tetrachloro-3,6-diphenylglycouril) was obtained from Pierce Chemical (Rockford, IL). Percoll and density marker beads were obtained from Pharmacia Biotech (Uppsala, Sweden). All other reagents were the purest grade available.
Radiolabeling.
Five microliters of DMSO solution containing DTPA (6 mg/ml) were added
to 250 µl of rhEPO solution (800 µg/ml) and allowed to stand for 30 min at room temperature. The resulting mixture was chromatographed on a
Sephadex G-25 PD-10 column (Pharmacia Biotech) with 50 mM
HEPES buffer (pH 7.0) containing 0.05% Tween 20 as eluent to obtain
fractions of DTPA-rhEPO. One hundred microliters of 0.2 M
citrate-acetate buffer (pH 5.0) and
111InCl3
(200 µCi) were added to 100 µl DTPA-rhEPO solution (150 µg/ml).
The resulting mixture was allowed to stand for 30 min in an ice bath
and then chromatographed on a Sephadex G-25 PD-10 column (Pharmacia
Biotech) by use of 50 mM HEPES buffer (pH 7.0) containing 0.05% Tween
20 as eluent to obtain fractions of
111In-rhEPO.
125I-rhEPO was prepared by the
Iodo-Gen method described previously (11). The specific radioactivity
of 111In-rhEPO and
125I-rhEPO was 4.5 and 17.6 µCi/µg, respectively, as determined by gel filtration assay. The
radiochemical purity of
111In-rhEPO and
125I-rhEPO was 98 and 97%,
respectively, as determined by gel filtration.
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Animals. Male Sprague-Dawley rats (JCL:SD, Clea Japan, Tokyo, Japan) were allowed to acclimatize to the laboratory environment for 1 wk; the experiment was started when they had reached 7 wk of age. Animal rooms were maintained at constant ambient temperature and a relative humidity of 24°C and 55%, respectively, throughout the experimental period. A standard rodent feed in pellet form (CE-2, Clea Japan) and tap water ad libitum were available throughout the study. This study was carried out in accordance with the Declaration of Helsinki and the Guide for the Care and Use of Laboratory Animals as adopted and promulgated by the US National Institutes of Health.
rhEPO treatment. rhEPO was administered intravenously at doses of 0, 1, 5, and 25 µg/kg, twice over a 4-day period, the first dose being given on day 0 and the second on day 2.
Preparation of cell suspension.
The spleens were disrupted by using a spatula to press them through a
stainless steel screen (100 mesh) into -MEM containing 20% FCS.
Bone marrow cells were scraped from the femurs and were suspended in
-MEM containing 20% FCS. The cells were centrifuged for 5 min at
400 g. The cell pellets were
resuspended in
-MEM containing 20% FCS.
Measurement of burst-forming unit erythroids and CFU-E. Numbers of burst-forming unit erythroids (BFU-E) and CFU-E were estimated using the methylcellulose method of Iscove and Sieber (6).
Binding of 125I-rhEPO to splenic cells.
Binding experiments were performed in test tubes at room temperature
(24°C) for 1 h in a total volume (1 ml) of -MEM
containing tracer amounts of
125I-rhEPO (~10,000 counts/min)
and 3 × 107 splenic
cells/ml, with or without unlabeled rhEPO. After incubation, the cells
were centrifuged at 400 g for 5 min.
The supernatant was aspirated, the cells were washed twice with 2 ml
ice-cold
-MEM, and the radioactivity of the final pellet was
counted. The specific binding was determined by subtracting the binding in the presence of 500 ng/ml unlabeled rhEPO from the total binding observed in the absence of unlabeled rhEPO.
Effect of rhEPO treatment on the migration of the cells labeled metabolically with 111In-rhEPO or [3H]thymidine. 111In-rhEPO (0.4 µg/kg) or [3H]thymidine (100 µCi/kg) was administered intravenously into the tail vein. rhEPO (25 µg/kg) was administered intravenously at both 12 h (day 0) and 60 h (day 3) after administration of 111In-rhEPO or [3H]thymidine. The rats were allowed to bleed to death after cardiac puncture under ether anesthesia on day 0, 1, 2, 3, or 4 after the first EPO treatment. The tissues were removed and weighed. Blood was transferred to heparinized tubes and centrifuged at 15,000 rpm for 3 min to obtain plasma. Then 1 ml of plasma and the entire tissue were subjected to gamma or scintillation counting, as will be described, to determine 111In and 3H radioactivity, respectively.
Separation of cells.
The splenic cells obtained from three rats were mixed. Two milliliters
of cell suspension were mixed with 43 ml of 45% Percoll in -MEM
containing 20% FCS, and the mixture was centrifuged at 10,000 g for 90 min. The temperature was
20°C. In a parallel tube, density marker beads (Pharmacia Biotech)
suspended in the same medium were processed in a similar fashion. After
the run, 2.5-ml fractions were collected from the bottom of the
gradient and diluted with 2.5 ml
-MEM. The cells were centrifuged at
400 g for 5 min, and the cell pellets
were resuspended in 2 ml of
-MEM and centrifuged at 400 g for 5 min. Finally, the cell pellets
were resuspended in
-MEM.
Distribution of 3H-labeled splenic
cells.
rhEPO (25 µg/kg) was administered intravenously to rats 12 h after an
intravenous administration of
[3H]thymidine (100 µCi/kg). The spleens were removed 2 days after rhEPO administration.
The cells in four fractions corresponding to the BFU-E density were
collected by the procedure just described. The cells were washed twice
with -MEM and resuspended in this solution. Untreated rats received
injections into the tail vein of 2 ml/kg of the isolated cell
suspension so obtained (20,000-30,000 dpm/head). The rats were
then allowed to bleed to death after cardiac puncture under ether
anesthesia 1, 4, or 24 h after dosing. The spleen, liver, lung, kidney,
and bone marrow were then removed.
Measurement of 3H radioactivity. The cells were solubilized with Soluene-350 (Packard Instrument, Meriden, CT). Entire tissues or portions of tissues were dried at room temperature for 1 wk and combusted in a sample oxidizer (Aloka, Tokyo, Japan). The radioactivity was recovered as 3H20 and measured by a scintillation counter (Beckman Instrument, Fullerton, CA).
Statistical methods. Comparisons of means were performed using one-way analysis of variance followed by Tukey's test. Statistical significance was taken as P < 0.05.
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RESULTS |
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Changes in target cell numbers and
125I-rhEPO specific binding in splenic cells
after repeated administration of rhEPO.
Both the numbers of CFU-E and BFU-E in the spleen and the
specific binding of 125I-rhEPO to
the splenic cells were measured after repeated administration of rhEPO
to rats. rhEPO treatment caused an increase in the number of CFU-E in
the spleen depending on the dose administered (Table 1). The number of
BFU-E did not increase after doses of 1 and 5 µg/kg but increased
only after a dose of 25 µg/kg (Table 1). The CLup in the spleen, estimated
by dividing the tissue concentration by the area under the plasma
concentration curve after intravenous administration of a trace amount
of 125I-rhEPO to rhEPO-treated
rats (7), also increased significantly in a dose-dependent manner
(Table 1) (7). The CLup by the spleen was almost proportional to the number of CFU-E in the spleen (Fig. 1A). No correlation was
observed between the number of BFU-E and
CLup by the spleen (Fig.
1B). To clarify the relationship between CFU-E and EPO receptor numbers, the specific binding of 125I-rhEPO to isolated splenic
cells was measured. Because this study was performed at a tracer
concentration (~0.2 pM), ~1/1,000 of the dissociation constant
(Kd) for EPO
receptors (180 pM) (1), this EPO-binding should represent EPO receptor
density on the surface of splenic cells if the affinity for the
receptor remains unchanged. As shown in Fig. 1, the increase in
CLup after EPO administration is
proportional to the increase in CFU-E numbers but not to that in BFU-E
numbers. Because the EPO receptor is known to be expressed almost
exclusively on CFU-E among all the splenic cells, this result suggests
that upregulation of CLup results
from the increase in either EPO receptor numbers on CFU-E or the number
of CFU-E in the spleen. In addition, the CFU-E numbers were also
proportional to the specific binding of
125I-rhEPO to all the splenic
cells (Fig. 1). This result suggests that the receptor number or
receptor binding activity per individual CFU-E is unchanged by EPO
treatment, whereas the number of CFU-E is increased after EPO
administration, causing upregulation of CLup.
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Migration of CFU-E after repeated administration of rhEPO.
We previously found that repeated administration of rhEPO caused an
increase in CLup by the spleen,
whereas the increase in CLup by
bone marrow was minimal (7). In the present study, we hypothesized that
such a difference in the degree of upregulation between bone marrow and
spleen might be due to migration of the target cells expressing EPO
receptors (mainly regarded as CFU-E) from bone marrow to spleen. In
Fig. 2, CFU-E-expressing EPO receptors were
labeled with 111In via EPO
receptor-mediated endocytosis in vivo by intravenous administration of
111In-rhEPO. The level of
111In radioactivity in bone marrow
fell without rhEPO treatment (Fig. 2A). After rhEPO treatment, this
reduction in 111In radioactivity
in bone marrow was significantly more marked (Fig.
2A). Thus rhEPO stimulates the
release of 111In-labeled cells
from bone marrow. rhEPO treatment also had a similar effect in the
spleen (Fig. 2B). This result
suggests that rhEPO does not stimulate migration of CFU-E-expressing
EPO receptors into the spleen. The radioactivity in blood cells
increased, with or without rhEPO treatment, suggesting that at least
some of the CFU-E and/or more mature cells were released into the blood
stream (Fig. 3). The levels of
radioactivity in other tissues were unchanged by rhEPO treatment,
showing that rhEPO had no effect on these tissues (Fig. 3).
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Migration of BFU-E into the spleen.
[3H]thymidine was
administered to rats to metabolically label growth cells. The level of
3H radioactivity in bone marrow in
both control and EPO-treated rats decreased from day
1 to day 4 (Fig.
4A). The
fall in radioactivity produced by EPO treatment (Fig.
4A) might be due to the maturation of cells and/or the release of cells from bone marrow. On other hand,
the level of 3H radioactivity in
spleen increased from day 1 to
day 2 (Fig. 4B). This result suggests that the
migration of a certain type of growth cell labeled with
[3H]thymidine into the
spleen was stimulated by rhEPO, predominantly from day
1 to day 2. After
day 2, the radioactivity in the spleen remained unchanged between days 2 and
3 (Fig.
4B), although the radioactivity in
bone marrow continued to fall (Fig.
4A). In considering the reason for
this phenomenon, it should be noted that
3H-labeled cells in spleen will be
released and/or degraded. Therefore, this result can be explained if
the influx rate of 3H-labeled
cells to the spleen equals the efflux rate of the cells from the
spleen. BFU-E is a type of growth cell and precursor of CFU-E. If such
[3H]thymidine-labeled
cells, whose migration into spleen was stimulated by rhEPO, include
BFU-E, and BFU-E migrating in this fashion differentiate to CFU-E in
the spleen, this would explain the difference in the degree of
upregulation of CLup between bone
marrow and spleen. To clarify whether the increase in radioactivity was
due to an increase in BFU-E, cell fractionation was performed using
Percoll to identify the BFU-E and CFU-E fractions in the
[3H]thymidine-labeled
splenic cells. To establish a cell-fractionation system, isolated bone
marrow cells in untreated rats were fractionated by centrifugation
through a Percoll density gradient. The number of CFU-E and BFU-E in
each fraction was measured by colony assay (Fig.
5). BFU-E and CFU-E from rat bone marrow
were eluted in fractions having a density of 1.050 and 1.065, respectively, indicating that BFU-E can be separated from CFU-E by the
difference in their density (Fig. 5). Then, the spleen was removed from
the rats treated with rhEPO after
[3H]thymidine
administration, and the splenic cells were fractionated by
centrifugation through this Percoll density gradient. The radioactivity in both the BFU-E and CFU-E fractions of the splenic cells increased from day 1 to day
2 after the first EPO treatment (Fig.
6). The radioactivity in the CFU-E fraction
increased further from day 2 to
day 3, whereas the radioactivity in
the BFU-E fraction fell (Fig. 6). The radioactivity in the CFU-E
fraction then fell from day 3 to
day 4. Thus we have demonstrated that
BFU-E migrate to the spleen and differentiate there into CFU-E. The
radioactivity in each fraction did not show a marked change without
rhEPO treatment (Fig. 6).
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Spleen-specific distribution of
[3H]thymidine-labeled BFU-E
fraction.
The
[3H]thymidine-labeled
splenic cells in the BFU-E fraction after centrifugation through a
Percoll density gradient were obtained from rats 2 days after rhEPO
treatment. The levels of radioactivity in tissues were measured after
intravenous administration of the cells thus obtained to control rats.
The radioactivity in lung and liver (37 and 30%, respectively) was
higher than in other tissues at 1 h after administration. Then these
levels fell with time (Fig. 7). However,
the radioactivity in spleen increased with time and accounted for
~18% of the dose, which was much higher than that in lung and liver
at 24 h after administration of the cells. No radioactivity could be
detected in bone marrow at any time. When the detection limit of this
assay is considered, the radioactivity in bone marrow should correspond
to <17% of the dose. These results suggest that BFU-E might move
into the spleen via blood flow and stay there.
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DISCUSSION |
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A single dose of rhEPO causes transient downregulation of CLup by both bone marrow and spleen, whereas repeated rhEPO administration results in marked upregulation, especially by the spleen (7). We believe that the upregulation of CLup by the spleen is due to an increase in CFU-E, because EPO receptors are predominantly expressed in CFU-E, and both the CLup by spleen and the specific binding of 125I-rhEPO to splenic cells were almost proportional to the number of CFU-E in the spleen (Fig. 1). This result suggests that the increased tissue uptake of rhEPO by the spleen is not due to an increased number of receptors per cell but to an increased number of CFU-E per whole spleen. A similar phenomenon has also been observed when endogenous EPO is increased in anemia. Akahane et al. (1) reported a good correlation between the number of CFU-E in the bone marrow of anemic rats and the specific binding of 125I-rhEPO to bone marrow cells isolated from such rats. This increase in target cells after repeated administration of rhEPO results in an increase in the tissue uptake of rhEPO, which then leads to an increase in the total body clearance of rhEPO. Therefore, such upregulation of CLup may be one of the homeostatic regulators that play a role in providing excess EPO to maintain the plasma EPO level constant. As far as G-CSF is concerned, similar upregulation of G-CSF receptors has also been observed. Kuwabara et al. (16) reported that the administration of the G-CSF derivative, nartograstim, caused transient downregulation of G-CSF receptors in both bone marrow and spleen, followed by marked upregulation in spleen. The authors suggested that such upregulation of G-CSF receptors in the spleen might be due to an increase in neutrophils expressing G-CSF receptors in the spleen (16). Thus the increase in the number of target cells leads to the upregulation of receptors also after administration of G-CSF.
We hypothesized that the difference in the degree of upregulation between bone marrow and spleen might be due to stimulation of the migration of CFU-E from bone marrow to spleen after rhEPO treatment. To examine this hypothesis, 111In-rhEPO was administered intravenously, and the radioactivity in each tissue was monitored (Figs. 2 and 3). EPO receptors are predominantly expressed in CFU-E, and 111In-rhEPO can be taken up into CFU-E by RME after intravenous administration of 111In-rhEPO. The 111In-DTPA moiety conjugated to the carrier protein has been reported to accumulate in the intracellular space by exchanging the DTPA moiety of the carrier protein with intracellular iron-binding protein (2). Its efflux from the cells was much slower than that of 125I radioactivity after the endocytosis of 125I-labeled protein produced by chloramine T. Therefore, proteins labeled with 111In can be used in the present approach involving the labeling of CFU-E-expressing EPO receptors with 111In via RME by use of 111In-rhEPO in vivo (Fig. 2). The ratio of the amount of radioactivity in bone marrow to that in spleen was ~20 at 12 h after administration of 111In-rhEPO (Fig. 2). This was comparable with the CLup ratio (~13) of bone marrow to that of spleen (8) assessed using 125I-rhEPO. Furthermore, the administration of excess (125 µg/kg) rhEPO inhibited the distribution of 111In-rhEPO both to bone marrow and spleen (data not shown). These results suggest that CFU-E was labeled with 111In-rhEPO via RME. The level of 111In radioactivity in bone marrow fell with time in control rats, whereas rhEPO treatment stimulated this reduction in radioactivity in bone marrow (Fig. 2). This result suggests that repeated rhEPO administration stimulates the release of CFU-E from bone marrow. If this released CFU-E migrates to the spleen, the 111In radioactivity in spleen should be increased by rhEPO treatment. However, the level of 111In radioactivity in the spleen also fell, and this reduction was significantly stimulated by rhEPO treatment (Fig. 2). This finding suggests that rhEPO stimulates the release of CFU-E from both tissues and refutes our hypothesis that the migration of CFU-E-expressing EPO receptors directly causes the upregulation of EPO receptors after rhEPO administration.
Our next hypothesis was that the increase of CFU-E in the spleen may result from the migration of its precursor, BFU-E, into the spleen, followed by differentiation there to CFU-E. When [3H]thymidine was administered in vivo to metabolically labeled growth cells including BFU-E, the level of radioactivity in bone marrow was significantly reduced by rhEPO treatment, whereas the level of radioactivity in the spleen was significantly increased (Fig. 4). Thus a certain type of the growth cell labeled with [3H]thymidine moved into the spleen from bone marrow after treatment with rhEPO. Nijhof et al. (22) found that, in mice, there is a change in the number of CFU-E and BFU-E in bone marrow and spleen after treatment with rhEPO, and they concluded that a redistribution of BFU-E from bone marrow into spleen occurs after rhEPO treatment. To prove the migration of BFU-E into the spleen and the subsequent differentiation of BFU-E to CFU-E there, we examined the kinetics of 3H-labeled BFU-E and CFU-E in the spleen (Fig. 6). Although the density of CFU-E and BFU-E in rats was unknown, the data on the purification of CFU-E in mice suggested that there was a difference in density between CFU-E and BFU-E (21), which prompted us to take advantage of this and to devise a separation method. Cell fractionation was performed on isolated splenic cells with use of Percoll in the present study, and BFU-E and CFU-E were found in fractions with densities of 1.050 and 1.065, respectively (Fig. 5). The density of BFU-E and CFU-E was similar to that of human BFU-E and mouse CFU-E, respectively (19, 21). The kinetics of 3H radioactivity in both BFU-E and CFU-E fractions demonstrate that BFU-E might move from bone marrow into spleen and differentiate there to CFU-E. To support our hypothesis, the [3H]thymidine-labeled splenic cells in the BFU-E fraction were isolated and administered to rats to clarify whether BFU-E moves specifically into spleen (Fig. 7). Initially, the radioactivity administered was mainly distributed to lung and liver (Fig. 7). The distribution of radioactivity to these organs may be relatively nonspecific, judging from the observation that this radioactivity fell over time (Fig. 7). On the other hand, the radioactivity in spleen increased with time and was much higher than that in lung and liver (Fig. 7). These results suggest that BFU-E can move into the spleen via blood flow and stay there. No radioactivity was detected in bone marrow at any time (Fig. 7). However, this result does not mean that BFU-E does not move into bone marrow. In the present study, the yield of 3H-labeled BFU-E was limited and, consequently, the amount of radioactivity administered was small. Therefore, the radioactivity in bone marrow may not be detectable even if a small portion is actually distributed. As far as the distribution of cells in the BFU-E fraction to lung and liver is concerned, one possible mechanism for such distribution is the nonspecific trapping by the capillary space. We have previously reported that cancer cells injected through the femoral and portal veins were predominantly distributed to the lung and liver, respectively, within 1 min. Such rapid distribution was ascribed to the phenomenon whereby the cells became clogged within the capillary because of their larger size (20).
In the present study, to clarify the disposition kinetics of target cells, radioactivity was monitored after growth cells were labeled by [3H]thymidine. It is possible that incorporated [3H]thymidine could be shed because of cell death and could be reutilized by cells other than those that are initially labeled and selected. In the experiment shown in Fig. 7, the radioactivity in the spleen 24 h after intravenous administration of [3H]thymidine was <1% of the administered dose (data not shown). This result suggests that the reutilized [3H]thymidine makes only a small contribution to the total radioactivity found in the spleen (Fig. 7). However, even when such an experiment was performed, the possibility that a small amount of reutilized 3H radioactivity might interfere with the experimental results cannot be completely neglected. In addition, in the experiments shown in Figs. 4 and 6, this possibility is undeniable. [3H]thymidine labeling of isolated cells is useful for following disposition kinetics in the body. Nevertheless, it is possible that reutilization of [3H]thymidine might be a limitation of this [3H]thymidine- labeling technique. More detailed experiments should be performed before rejecting this possibility.
We can speculate on the mechanism of the specific residence of BFU-E in the spleen from the results of our previous report showing that no upregulation of the CLup of 125I-rhEPO by the spleen was found after repeated rhEPO administration in W/Wv mice, which have a mutation of the c-kit (stem cell factor receptor) gene (4). This result suggests that c-kit is essential for the upregulation of CLup by the spleen. The stem cell factor is a membrane-binding protein in stromal cells that is involved in maintaining the microenvironment in hematopoietic tissues (4). The binding of c-kit on hematopoietic bone marrow cells to the stem cell factor activates integrin on the cell surface, causing the cells to bind strongly to fibronectin (3, 13), which is expressed in the extracelluler matrix of the microenviroment of hematopoietic tissues. Therefore, it can be speculated that this binding of c-kit to the stem cell factor may result in the increased stability of these hematopoietic cells in the microenvironment of hematopoietic tissues. The integrin of those cells in W/Wv mice cannot be activated because of the mutation of c-kit in W/Wv mice (10). Therefore, the hematopoietic cells in W/Wv mice are unable to bind to fibronectin. Accordingly, the migration of BFU-E into the spleen may come from binding to the stem cell factor in splenic stromal cells, this binding resulting in the activation of integrin in BFU-E. The BFU-E in W/Wv mice, therefore, cannot remain in the spleen because of inactivation of the integrin.
The BFU-E increase in spleen after rhEPO administration might arise from bone marrow, judging from the observation that the [3H]thymidine radioactivity in bone marrow is significantly reduced by rhEPO (Fig. 6) and that BFU-E are known to be almost exclusively confined to bone marrow, spleen, and blood. There are two possible mechanisms for the increase in the number of BFU-E in spleen: 1) the release of BFU-E from bone marrow into the bloodstream is stimulated, and 2) the adhesion of BFU-E from the bloodstream to the spleen is stimulated after rhEPO administration. The latter possibility is unlikely, because the distribution of isolated 3H-labeled cells in the BFU-E fraction to the spleen found in Fig. 7 was not stimulated by rhEPO administration (unpublished data), and so the former is more likely. A recent study suggests that expression of c-kit might be involved in the release of BFU-E from bone marrow, because the expression of c-kit on BFU-E in peripheral blood was lower than that on BFU-E in bone marrow (26). This suggestion is comparable with our previous finding (8) that no upregulation of CLup by the spleen was observed in W/Wv mice. Thus c-kit may be involved in the release of BFU-E from bone marrow. However, the exact molecular mechanism for the stimulatory effect of EPO on this release of BFU-E needs to be clarified by further research.
As we have discussed, once 111In-labeled protein is taken up by the cells via RME, 111In radioactivity remains inside the cells (2). If we take advantage of this property, proteins labeled with 111In can be used to precisely estimate CLup (5). Pharmacokinetic theory indicates that the ratio of the 111In radioactivity in each tissue to the administered dose should be proportional to the ratio of CLup to total body clearance. Such a ratio represents the contribution of each organ to the overall elimination of the protein. rhEPO is eliminated by both saturable and nonsaturable clearance mechanisms (7). We have reported that the mechanism of saturable clearance is RME by bone marrow and spleen (7) and that the kidney might, at least partially, contribute to the nonsaturable clearance (7, 12). However, the contribution of other tissues to the nonsaturable clearance is still unknown. The amount of radioactivity in muscle and skin accounted for 15 and 9% of the dose at 36 h after intravenous administration of 111In-rhEPO. This distribution was very close to that in bone marrow (12% of the dose). This high distribution of 111In-rhEPO in muscle and skin suggests that these tissues may play a role as major clearance sites. The amount of radioactivity in muscle and skin was only 0.8 and 6% of the dose at 24 h after intravenous administration of 125I-rhEPO (28). 125I radioactivity is known to undergo easy efflux from cells after endocytosis and subsequent degradation of protein labeled with 125I. Therefore, most of the 125I-rhEPO taken up by muscle and skin should be metabolized at 24 h. The bioavailability (ratio of subcutaneous to intravenous area under the curve) of 125I-rhEPO after subcutaneous administration was ~0.5 (9), which also suggests metabolism of 125I-rhEPO by muscle and skin.
In conclusion, upregulation of the CLup of rhEPO by the spleen after EPO treatment might be due to migration of BFU-E from bone marrow to spleen and subsequent differentiation into CFU-E in the spleen. This is the first demonstration that the upregulation of cytokine receptors is due to stimulation of target cell migration by cytokines.
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ACKNOWLEDGEMENTS |
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We thank Chugai Pharmaceutical Company for providing rhEPO.
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FOOTNOTES |
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This study was supported in part by a Grant-in-Aid for Scientific Research provided by the Ministry of Education, Science and Culture of Japan.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for correspondence and reprint requests: Y. Sugiyama, Dept. of Biopharmaceutics, Graduate School of Pharmaceutical Sciences, Univ. of Tokyo, 7-3-1, Hongo, Bunkyo-ku, Tokyo 113-0033, Japan (E-mail: sugiyama{at}seizai.f.u.-tokyo.ac.jp).
Received 24 August 1998; accepted in final form 4 February 1999.
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