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Blood, Vol. 91 No. 4 (February 15), 1998:
pp. 1280-1287
No Neuronal Regulation of Murine Bone Marrow Function
By
Haakon B. Benestad,
Inger Strøm-Gundersen,
Per Ole Iversen,
Egil Haug, and
Arild Njå
From the Departments of Physiology and Neurophysiology, Institute of
Basic Medical Sciences, The University of Oslo, and The Hormone
Laboratory, Aker Hospital, Oslo, Norway.
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ABSTRACT |
Bone marrow is innervated by efferent (sympathetic) and afferent
nerves, but it is not clear whether these nerves affect cell formation
or release in any significant way. To elucidate this problem, we
studied mice neonatally sympathectomized with 6-hydroxydopamine and
adult mice in which one hind limb was surgically denervated. Progenitor
and transit cell numbers and proliferative activity were estimated in
bone marrow, blood, and spleen. In addition, we performed unilateral
electrical stimulation of nerve fibers to tibial marrow and applied a
cell mobilizing stimulus (bleeding, granulocyte colony-stimulating
factor injection, or intraperitoneal injection of a chemotactic
substance) to investigate cell egress from the marrow. Blood flow to
hindleg bone marrow was assessed with the radioactive microsphere
technique. Except for a smaller bone marrow cell population and lower
body weight in neonatally sympathectomized mice, we found no clear
indications that bone marrow innervation influenced cell production.
Also, the innervation did not detectably affect cell release from the
marrow. Electrical stimulation of hind limb nerves did not change the
blood flow to the marrow, whereas it markedly decreased blood flow to
the overlying muscle. We therefore conclude that no obvious function can be ascribed to tibial marrow innervation in the mouse.
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INTRODUCTION |
SYSTEMIC SIGNALS that have regulating
influence on the bone marrow function, ie, formation and release of
blood cells, are generally believed to be blood-borne (as reviewed by
Benestad and Laerum1). However, a regulatory role has been
suggested for the bone marrow innervation as well. For example, it has
been suggested that the fine regulation of proliferation and release of
hematopoietic cells is performed via nerve fibers that may be sensitive
to minute pressure changes (pressure from hematopoietic growth or from
blood flow), particularly in an organ that is protected from external
pressure by a rigid bony capsule.2
It has been shown that afferent and efferent sympathetic nerves lie
close to blood vessels within the marrow. Stromal and parenchymal cells
are also innervated, although to a lesser extent than the
vasculature.3-7 In addition to noradrenergic sympathetic nerve fibers, various peptidergic nerves to the bone marrow of rats and
guinea pigs have been identified,8 in particular fibers containing substance P (SP) immunoreactivity.9 SP and
neurokinin-A (NK-A), both coded by the preprotachykinin I gene, have
been reported to release stimulatory and inhibitory hematopoietic
cytokines from human bone marrow cells10-13 and blood
monocytes.14 Furthermore, SP is a costimulant of various
colony-stimulating factors10,15 and may even alone
stimulate the formation of erythrocytic and granulocytic
colonies.10 Finally, SP might affect the motility of
granulocytes and monocytes16 and granulocyte infiltration into mouse skin17 and might thus possibly increase the rate of cell release from the bone marrow. Classical autonomic nerve transmitters might also modulate hematopoiesis in situ.
Byron18 demonstrated that spleen colony-forming cells
initiate DNA synthesis after stimulation with  -adrenergic or
cholinergic agonists, and other workers19,20 have found a
cholinergic enhancement of megakaryocytopoiesis and granulocytopoiesis
in culture. On the other hand, Maestroni et al21,22 showed
that chemical sympathectomy of adult mice and
 1-adrenergic antagonists can enhance myelopoiesis and
platelet formation.
Further evidence of a neuronal regulation of bone marrow function was
provided by Foa,23 who claimed that electrical stimulation of the sympathetic chain or nerves sending branches to bone marrow led
to vascular contraction and mobilization of erythrocytes, granulocytes,
and immature cells into the blood of dogs. Similar findings have been
made in rats, in which reticulocytes and granulocytes were mobilized by
1.5 to 2 hours of stimulation of both sympathetic trunks, with or
without prior adrenalectomy or nephrectomy/ligation of renal
vessels.24,25
Nevertheless, firm evidence for neuronal regulation of bone marrow
function has to our knowledge not been presented. Therefore, we have
used both well-established and new approaches to determine whether
nerves can affect cell formation in or cell mobilization from bone
marrow of the mouse. Mice were treated with 6-hydroxy-dopamine (6-OH-DA) shortly after birth to destroy the sympathetic nervous system. Some of these mice were adrenalectomized to remove this additional source of catecholamines, which might simulate sympathetic nerve transmitters. Hematopoiesis and marrow cell release were compared
in these and sham-treated mice from the same litters. Furthermore,
denervated or nerve-stimulated tibial marrow was sampled and compared
with the contralateral control marrow in hormonally intact and in
adrenalectomized mice. Steady-state and regenerating bone marrows were
investigated. In the cell release studies, we examined whether nerve
stimulation could mobilize cells to the blood or affect mobilization
induced by injection of recombinant granulocyte colony-stimulating
factor (G-CSF) or a chemotactic agent.
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MATERIALS AND METHODS |
Mice.
Female NMRI/Bom or ICR/OlaH mice were used. They were at least 4 and
usually 7 to 15 weeks old and had free access to pellet food and tap
water. A few experiments were performed with the youngest mice to find
out whether nerves could play another role in a developing than in a
mature hematopoietic system.
Except for the cell release experiments (see below), we performed
surgery under midazolam (Dormicum; Roche, Basel,
Switzerland) plus fentanyl/fluanisone (Hypnorm; Janssen, Beerse,
Belgium) anesthesia and, except for the blood flow
determinations (see below), the mice were killed with carbon dioxide
gas.
Four main experimental designs were used: (1) cell generation in
chemically sympathectomized mice; (2) cell generation in surgically
denervated tibial marrow; (3) cell release in chemically sympathectomized mice; and (4) cell release from tibia during electrical nerve stimulation.
All experimental protocols were approved by the regional animal
experimentation committee.
Chemical sympathectomy.
To destroy sympathetic neurons, mice received 6-OH-DA (Sigma Chemical
Co, St Louis, MO) by subcutaneous injection (4 mg/mL in saline; 25 µL
per gram of mouse weight; 4 injections; one every second day from when
they were 2 days old)26 and otherwise treated as
described.27 Control littermates received solvent
injections.
Irides from 6-OH-DA- and solvent-treated mice were examined to
establish the completeness of the chemical sympathectomy. They were
stained with glyoxylic acid28 to ascertain that the
denervation procedure had effectively removed all the catecholaminergic
innervation. In only a few cases were insignificant remnants of nerve
fibers found.
Surgical denervation.
One of the sciatic nerves was cut at the level of the sciatic notch. In
initial experiments the femoral nerve was also cut just below the
inguinal ligament. At the same time we performed a contralateral sham
operation.
Smears made from marrow plugs, washed out of the tibiae with jets of
medium, and small arteries sampled close to the tibial bones were
stained with glyoxylic acid to check the denervation of the tibial
marrow. Preparations from normal or sham operated legs showed a
prominent plexus of fluorescent nerve fibers that was largely confined
to the blood vessels (Fig 1). Two to 3 days after sciatic nerve section, this system of nerve fibers was either completely absent or reduced to occasional patches. This indicates that
the sympathetic fibers to the tibial marrow are carried almost exclusively by the sciatic nerve (see also Weiss and
Root29).
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| Fig 1.
Sympathetic nerve fibers in tibial bone marrow. A dense
plexus of nerve fibers with brightly stained varicosities is present around blood vessels. Digital image of normal bone marrow removed from
mouse tibia and stained with glyoxylic acid. The diameter of the large
vessel is about 30 µm. 40× water immersion objective.
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Electrical stimulation.
To study stimulation-induced cell release, mice were deeply
anesthetized with chloral hydrate (0.2 g/kg) and pentobarbitone (0.05 g/kg) intraperitoneally (IP) and mechanically ventilated. An additional dose of the anesthetic mixture was supplied every 30 minutes. Both the left and the right sciatic nerves were cut, and the
distal cut end of the right nerve was stimulated electrically. After a
single test stimulus, further muscle contractions were eliminated by a
neuromuscular blocking agent (1 mg/kg IP Alloferin; Hoffmann-LaRoche & Co AG, Basel, Switzerland). The nerve was then stimulated at 5 Hz
continuously or 20 Hz for 5 seconds every 20 seconds for 50 minutes (so
that the total number of stimulus pulses was 15,000 in
both cases), with symmetrical bipolar current pulses (± 1 mA, 0.5 milliseconds each). The 5 Hz continuous stimulation was intended to
release only adrenergic transmitters, whereas the 20 Hz pattern was
intended to also release neuropeptides (see Lundberg et
al30). The animals were then killed with neck luxation and
tissues were removed immediately for analysis.
Adrenalectomy.
Some mice were delivered adrenalectomized or sham adrenalectomized from
the vendor (Gml. Bomholtgaard, Ry, Denmark) or the national importer of
experimental animals (SIFF, Oslo, Norway). These procedures were later
performed in our own laboratory.27 These mice drank 0.9%
saline. The adrenalectomy had been performed at least 4 weeks before
the cell generation experiments and at least 4 days before the cell
release experiments.
To monitor the completeness of the adrenalectomies, corticosterone (the
major adrenal glucocorticoid hormone in the mouse) in slightly diluted
mouse heparin plasma was analyzed after ether extraction of 100 µL
plasma with a previously described radioimmunoassay.27 The
intra-assay coefficient of variation is between 6% and 10%. In 8 hormonally intact mice, the quartile interval for the corticosterone concentration was 350 to 570 nmol/L (for the plasma protein
concentration, 28 to 32 mg/mL). The corresponding data for 23 adrenalectomized mice were 80 to 170 nmol/L and 28 to 35 mg/mL,
respectively. Three mice in this group had values indicating incomplete
removal of the adrenals, which did not significantly affect the median
values.
Bone marrow, spleen, and blood examinations.
The surgically denervated tibial bones or the femurs from chemically
sympathectomized mice were dissected free and crushed in a mortar with
cold serum-containing culture medium. The marrow cells were suspended
by vigorous pipetting, counted with a Coulter counter, smeared,
stained, and classified microscopically (200 cells counted from each
mouse, the identity of the preparation being unknown to the examiner).
Megakaryocytes were scored after acetylcholine esterase staining.
The differential counting of hemoglobin-containing nucleated cells was
facilitated by prestaining the smears with o-dianisidine.31
Reticulocytes were stained with Brilliant cresyl blue and 500 to 1,000 cells were scored from each sample of mouse blood. Total and
differential blood cell counts (100 to 200 cells per mouse) were
performed as for the marrow. Alternatively, reticulocyte and
differential counting was performed with automatic cell counters (R-1000 Sysmec [Kobe, Japan] and Technicon H-1 [Tarrytown, NY]).
Standard methods were used to make single-cell suspensions of spleen
cells, to determine the packed blood cell volumes (hematocrit), and to
analyze plasma protein concentrations.
Regenerating marrow cells were obtained 3 days after injection of 200 mg/kg IP cyclophosphamide (Sendoxan; Asta Medica, Frankfurt, Germany).
We enumerated granulocyte-macrophage colony-forming cells (G/M-CFC = GM-CFC, G-CFC, and M-CFC) by culturing 5 × 104
steady-state or 2 × 104 regenerating marrow cells and
0.5 × 106 or 1 × 106 spleen cells
in methylcellulose-containing medium (1 mL) for 7 days with pokeweed
mitogen-stimulated spleen cell conditioned medium as a source of
colony-stimulating factors. Cell aggregates with more than
approximately 50 cells were scored as colonies.32
As an alternative to the standard colony scoring method, dishes were
scored automatically with a digital image analysis system applied to
video pictures of the dishes.33 Viable cells in the cultures had then been stained supravitally beforehand.34
Flow cytometry of single-cell suspensions of bone marrow was performed
with a FACScan (Becton Dickinson, Mountain View, CA), the unstained
cells (10,000 cells per sample) being classified according to their
light forward and side scatter properties. The positions of mature
granulocytes, small lymphocyte-like cells, and macrophage-like cells in
the dot plots thus obtained had been determined in pilot experiments
with less heterogeneous cell populations. These were peritoneal cells
harvested at different intervals after injection of a chemotactic agent
(Bacto-Tryptone; see below) and subjected to both flow cytometry and
standard differential counting.
Bone marrow proliferative activity was assessed with measurements of
3H-thymidine incorporation 1 hour after the intravenous
(IV) injection of 37 kBq 3H-thymidine/g body weight (code
TRK 120; Amersham, Amersham, UK) in approximately 0.2 mL
medium. Radioactivity in trichloroacetic acid cellular precipitates and
in a diluted sample of the 3H-thymidine prepared for
injection was counted in a -counter.
Granulocyte mobilizing agents.
Bacto-Tryptone (10% in saline, 1 mL per mouse; Difco Labs, Detroit,
MI), which is a caseïn digest, was used to mobilize bone marrow
neutrophils, because it has a strong, chemotactic effect after IP
injection and also markedly increases the concentration of these cells
in the blood.35 Human recombinant G-CSF (5 µg/kg IV;
Chugai Pharmaceutical Co, Ltd, Tokyo, Japan) was also used to release
marrow neutrophils.36
To assess the potency of G-CSF to mobilize PMN to blood, we sampled
twice from a cut metatarsal vein; otherwise, blood was withdrawn from
the inferior caval vein of the killed mice.
Blood flow measurements.
We used the radioactive microsphere method to determine organ perfusion
in anesthetized mice. This method has been thoroughly described by
Iversen et al,37 as used in the present study and also
validated for the mouse.38 Briefly, about 40,000 well-mixed microspheres (diameter, 16 µm; NEN, Boston, MA) labeled with either 153Gd or 51Cr were injected as a bolus into the
left ventricle after the chest had been opened in the midline. The two
femurs, tibiae, and the two soleus muscles were removed, as well as the
two kidneys. The radioactivity of these samples was determined in an
Auto-Gamma 5220 counter (Packard, Downers Grove, IL). The
lowest number of microspheres deposited in any sample exceeded 250.
Statistics.
The values are expressed as medians with or without their corresponding
95% confidence intervals, determined with a nonparametric method
(program MINITAB; Minitab Inc, State College, PA). If the median of a
control group was not included in the 95% confidence interval of the
test group, and vice versa, then the two groups of data were considered
significantly different at the 5% level. With only one of the group
medians falling outside the other group's interval, the two-sided
significance was tested with Wilcoxon-Mann-Whitney two-sample test
(MINITAB). When appropriate, Wilcoxon's test for paired comparisons or
the nonparametric Wilcoxon-van Elteren test for paired groups of
data39 was applied.
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RESULTS |
Bone marrow hematopoiesis.
To examine the effect of the sympathetic innervation on hematopoiesis,
we studied mice that had been chemically sympathectomized by 6-OH-DA
shortly after birth. We found no significant influence of this
treatment on blood cell concentrations (Fig
2). However, the bone marrow cellularity was lower in 6-OH-DA mice than
in the controls, whereas the 3H-thymidine incorporation per
femur and the differential counts were apparently normal
(Fig 3). No differences were detected
between tests and controls concerning the concentrations of granulocyte and macrophage colony-forming cells, either in steady-state or in
regenerating bone marrow and spleen (Table
1). However, the cellularities of marrow and spleen were lower in the
sympathectomized mice 3 days after cyclophosphamide had been
administered to kill cycling cells and provoke a regenerative response
(Table 1). The body weights were also lowest in the experimental group
(95% confidence intervals of the medians: males, 33 to 39 g v
37 to 42 g; females, 26 to 30 g v 27 to 32 g;
P = .001 for all litters analyzed according to treatment group
and gender).
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| Fig 2.
Blood cell concentrations in chemically sympathectomized
and control animals. Median values with their 95% confidence intervals and the number of mice examined (top columns) are given.
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| Fig 3.
Decreased femoral marrow cellularity, but no change in
proliferative activity and differential counts in chemically
sympathectomized mice. The mice were killed 2 hours after
Bacto-Tryptone injection. *P = .002. (For these data pooled
with data from another series of experiments [17 + 19 cyclophosphamide-treated mice], P < .0001). Median values
with their 95% confidence intervals and the number of mice examined
(top columns) are given.
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Table 1.
Effects of Chemical Sympathectomy on Cell Numbers in
Femoral Bone Marrow and Spleen During Steady-State and During
Regeneration After Cyclophosphamide IP
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We then let each experimental animal serve as its own control by
surgically denervating one hind limb. Paired comparisons did not show
significant differences between the denervated and the intact tibia.
This applies to numbers of both progenitors (G/M-CFC) and more mature
cells, 2 or 7 days after one-sided denervation of normal as well as
adrenalectomized mice (Table 2). Similarly, experiments with adrenalectomized or hormonally intact mice did not
disclose any consistent effects of denervation on regenerating marrow
examined 3 to 6 days after the injection of cyclophosphamide (eg, for
all days taken together, the 95% confidence interval for the
difference in cellularity between innervated and denervated tibiae was
[ 1.1, 0.1] × 106 cells, P = .08, n = 29 mice). A representative experiment is shown in
Fig 4. The regeneration model had been
validated in experiments where we measured tibial marrow cellularities
before and 3, 5, and 6 days after cyclophosphamide. Rapid cell
proliferation took place, with significant cellularity increases
between successive days of regeneration and between day 6 and the
steady-state level (data not shown).
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| Fig 4.
Bone marrow regeneration after cyclophosphamide
treatment. Eight mice had one leg denervated and the other sham
operated; thereafter, they received cyclophosphamide at 200 mg/kg IP
(day 0). From day 3 on, 2 mice were killed at each time point and
cellularities compared between innervated and denervated tibia. The
stippled belt depicts the 95% confidence interval for total tibial
cell number in untreated mice. Digital image analysis of the
methylcellulose dishes gave two sets of data in addition to the colony
numbers (G/M-CFC/tibia), ie, (1) the total cellularity per dish when 5 × 104 cells had been cultured for 7 days (ie, G/M
cytopoietic capacity per tibia in arbitrary units [au]) and (2)
median colony size. ( ) Innervated; ( ) denervated.
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The preparatory operations per se did not affect the hematopoietic
system as far as we could detect by enumerating blood, spleen, and bone
marrow cells 2 and 4 days after sham dener- vation and at least 4 weeks after adrenalectomy (data not shown).
Cell release from bone marrow.
We counted neutrophilic granulocytes (PMN) and reticulocytes. PMN is
the only mature cell type with a large storage compartment in the bone
marrow, and reticulocytes may also be rapidly released (mobilized).
Various humoral stimuli can release cells from the bone marrow. Our
mobilizing agent was Bacto-Tryptone injected IP 2 hours before cell
sampling in combination with bleeding approximately 10% of estimated
blood volume.
Neither the numbers of PMN in the peritoneal cavity nor the
concentration of reticulocytes in the blood was detectably different in
chemically sympathectomized mice and intact littermate mice (Fig 5). Moreover, we did not find
significantly different sizes of the neutrophil storage compartments in
the bone marrow after cell release in the two groups of mice (data not
shown).
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| Fig 5.
No significant effects on peritoneal and blood cell
populations of Bacto-Tryptone IP and slight (0.2 to 0.3 mL) bleeding. Blood reticulocyte change refers to reticulocyte percentages in venous
blood 2 hours after and just before the IP Bacto-Tryptone injection.
Median values with their 95% confidence intervals and the number of
mice examined (top columns) are given.
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Similarly, we could not detect any significantly different neutrophil
storage pool in the denervated compared with the innervated tibial
marrow in mice treated with Bacto-Tryptone
(Table 3).
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Table 3.
No Significantly Different Cell Numbers in
Denervated and Innervated Tibial Marrow After Granulocyte
Mobilization With Bacto-Tryptone
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In further experiments, the sciatic nerve was cut bilaterally and the
distal cut end was stimulated electrically for about 50 minutes on one
side. A mobilizing stimulus (Bacto-Tryptone IP or G-CSF IV) was applied
here in some of the experiments 3 to 4 hours beforehand. Again, we
could not detect any systematic differences in mature neutrophil
numbers between the two tibiae (95% confidence intervals for the
differences: [1.17, 0.02] × 106 cells with
mobilization and [1.62, 0.55] × 106 cells
without; the positive control being the presence of a blood granulocytosis after G-CSF treatment). Moreover, the nerves seemed to
have no influence on marrow blood flow, as expected,37
whereas a markedly different blood flow to the musculature of the two legs was found (Fig 6).
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| Fig 6.
No significant difference in blood flow was found between
denervated and stimulated tibial marrow, in contrast to overlying muscle. The ratio is between the right and left side in mice with denervated left and (neurally) stimulated right hindleg. Median values
are given with their 95% confidence intervals (except for the muscle,
in which the range is depicted) as well as the number of mice
examined.
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DISCUSSION |
Except for a modestly decreased total number of cells in the bone
marrow of neonatally sympathectomized mice, we found no conspicuous
nerve effects on cell generation in, or mobilization from, the marrow.
Our results are consistent with those from previous rat experiments,
where bone marrow nerves did not affect the energy metabolism of marrow
cells.40
A convincing demonstration of the roles of bone marrow nerves should
establish a relationship between nerve activity and cell formation in
or cell egress from the marrow. However, detailed, putative mechanisms
have been clarified, mostly in vitro, whereas the physiologic evidence
is still scant. For example, spleen colony-forming stem cells initiate
DNA synthesis in vitro after stimulation with -adrenergic
agonists,18 and human hematopoietic cells can be stimulated
by SP in vitro, apparently both directly and indirectly via stromal
cell release of cytokines.10-12 An elaborate, bidirectional interplay between bone marrow nerves on the one hand and stroma and
mononuclear cells on the other might exist, but has not been directly
shown, in which SP and cytokines through transcriptional mechanisms
might influence each other, both concerning secretion rates and
receptor expression. Therefore, although nervous regulation is
generally used for rapid adaptations (within a time frame of seconds,
and cell generation and mobilization from the marrow are outside this
range), it is plausible but not substantiated that the nerves to bone
marrow may play a regulatory role.
Indeed, a rich nervous network has been found in the marrow, consisting
of afferent fibers, mostly myelinated (4%), and efferent, sympathetic
fibers that are unmyelinated (96%), as reviewed by Yamazaki and
Allen.6 These investigators found that most efferent fibers
end close to marrow arterioles, but rarely in hematopoietic parenchyma
(5.3% of the fibers) or on venous sinus walls (2.7%). Their electron
microscopy and morphometric studies also showed what might be a
functional unit for signal conduction (the neuro-reticular complex)
composed of efferent nerves apparently synapsing with stromal cells
that are connected by gap junctions.
DePace and Webber25 described adrenergic nerves in rat bone
marrow, and Felten et al9 claimed that noradrenergic
postganglionic sympathetic nerve fibers innervate bone marrow and that
SP-immunoreactive fibers had been observed in some sections of bone
marrow. Weihe et al8 stated that, despite unsatisfactory
staining quality, they had observed nerve fibers containing calcitonin
gene-related peptide- and tachykinin-immunoreactivity in vascular and
nonvascular locations of rat and guinea pig bone marrow. Some fibers
with immunoreactivity to tyrosine hydroxylase and neuropeptide Y were also seen. However, it should be noted that non-nervous sources of the
most studied neuropeptide, SP, have been described, namely macrophages41,42 and endothelial cells,43,44
with both cell types being constituents of bone marrow stroma.
The picture is further complicated by the recent finding that NK-A,
formed by alternative splicing of the messenger of the preprotachykinin
1 gene (which also codes for SP), stimulates erythroid progenitors, but
inhibits proliferation of GM-CFC, partly through stromal secretion of
the inhibitory cytokines transforming growth factor- and macrophage
inflammatory protein-1.13 A neural fine-tuning of
hematopoiesis, based on these stimulatory and inhibitory SP and NK-A
mechanisms, might be consistent with our own finding that denervation
did not affect hematopoiesis, except that both SP and NK-A apparently
stimulated erythropoiesis in vitro.10-12
Maestroni et al21,22 found that adrenergic agents could
inhibit myelopoiesis and platelet formation in mice, both under steady-state conditions and after irradiation and syngeneic marrow transplantation. Based on experiments with chemically sympathectomized adult mice (by 6-OH-DA) and the 1-adrenergic antagonist
prazosin, they stated that the production of granulocytes and
macrophages seems to be under an inhibitory noradrenergic tone. Our
results do not support this view. It is possible that effects, which
may be both nonspecific and not related to the innervation of bone marrow, may take place after treatment of adult, nonadrenalectomized animals with 6-OH-DA. The same kind of objection could be raised against the interpretation of the prazosin results. On the other hand,
it could be argued that we had missed important clues by working with
neonatally sympathectomized mice. Thus, neonatally sympathectomized
rats had adapted to the small immune deviations observed shortly after
the 6-OH-DA treatment, when they were 42 to 56 days old, possibly due
to partial reinnervation of lymphoid organs and compensation by the
adrenal glands and other hormonal systems.45 However, nerve
regeneration was usually not detected in our experiments (iris
examinations; see the Materials and Methods), and we could not find
qualitatively different effects of sympathectomy or surgical
denervation between mice with and without adrenal glands.
Direct neural evidence for regulation of bone marrow function (obtained
by denervation, nerve stimulation, or measurement of nerve activity)
has, to our knowledge, been reported from only two laboratories.
Foa23 worked with anesthetized dogs, in which he cut the
distal tibia and recorded expansion and contraction of the tibial
marrow. Moreover, venous blood from the marrow was sampled. The sciatic
nerve and the lumbar sympathetic chain were cut and the distal ends
were stimulated. An immediate reduction in volume of the marrow was
found (see also Weiss and Root29). After the electrical
stimulation, the venous effluent from the marrow contained an increased
number of white and red blood cells, including immature cells. Because
the tibial marrow of young dogs is not very active hematopoietically,
experiments were also performed on rib marrow, with stimulation of
intercostal nerves and blood sampling from an intercostal vein. Here
too, the cell concentrations increased during the stimulation,
especially concerning the immature cells. However, there were no
sham-stimulated controls, no endotoxin or corticosteroid analyses, and
no plausible explanation of the mobilization of immature cells, which
are very scarce in the blood under normal physiologic circumstances.
Webber et al24,25 stimulated the sympathetic trunk
bilaterally in albino rats and observed release of reticulocytes and neutrophils into the blood. In some rats, adrenalectomy and nephrectomy were performed before the stimulation. However, absolute cell counts
were not performed only differentials. It is puzzling that no changes
were recorded in the controls; after all, they had been exposed to
laparotomy and some of them also to operations on the distal femurs (a
hole was drilled to aspirate marrow before the electrical stimulation).
A shift of lymphocytes to bone marrow46 and mobilization of
neutrophils to blood (as part of a corticoid stress response; reviewed
by Benestad Laerum1) might have explained their findings.
Moreover, based on data published by others,47-49 the
decline in marrow neutrophil percentage (loss of about 10% of all
marrow cells) should have led to a much more marked increase in blood
neutrophils than was actually found (about 28%) if the only relevant
perturbation of homeostasis in these rats was the stimulated marrow
cell egress. We have tried to reproduce the findings of Webber et al.
No significant reticulocytosis was observed, and no consistently
different blood PMN response between the lumbar sympathetic
trunk-stimulated and sham-stimulated rats (our unpublished observations) was observed either.
It has been reported, based on perfusion experiments, that increased
cell release accompanies increased blood flow through the bone
marrow.50 We hypothesized that perturbation of the sympathetic innervation of marrow blood vessels would lead to a change
in blood flow and hence a change in cell release rate. However, even
though the vasculature in the overlying muscle was sensitive to the
nerve stimulation, the marrow vasculature proved nonresponsive. This is
in accord with our previous findings in the rat with both
surgically37,51 and chemically52
sympathectomized animals. Similarly, stimulation of the sympathetic
trunk in the rat decreased blood flow to hind limb skin and muscle, but
not to the bone marrow (our unpublished observations).
Even though intricate nerve arrangements have been described in the
bone marrow (see above), sympathetic innervation may be without
physiologic significance, at least in the mouse (but possibly not in
other species, see Drinker and Drinker53). Conceivably, the
innervation may play a role under special, but unknown, conditions. For
example, studies that have used radioactive microspheres to measure
cerebral blood flow have shown that neural effects are minimal, except during extreme hypertension.54 The role of the
vasoconstrictor innervation may here be to protect the blood-brain
barrier against disruption should arterial pressure increase suddenly.
This teleologic explanation may not be valid for the bone marrow;
moreover, the generally accepted regulatory mechanisms for
hematopoiesis and cell release from the marrow are all humoral, namely
endocrine, paracrine, or autocrine.1,55
Our conclusion that tibial bone marrow innervation in the mouse does
not influence cell production or cell release (or at least not
directly, in a biologically significant way, and under ordinary
circumstances) supports the physiologic relevance of ex vivo
experiments on bone marrow.
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FOOTNOTES |
Submitted August 8, 1997;
accepted October 17, 1997.
Supported by grants from the Norwegian Research Council and from Anders
Jahre's Foundation for the Promotion of Science.
Address reprint requests to Haakon B. Benestad, MD,
Department of Physiology, Institute of Basic Medical Sciences,
University of Oslo, PO Box 1103 Blindern, N-0317 Oslo, Norway.
The publication costs of this article were defrayed in part by page
charge payment. This article must therefore be hereby marked
"advertisement" in accordance with 18 U.S.C. section 1734 solely
to indicate this fact.
| |
ACKNOWLEDGMENT |
The authors are indebted to Prof K. Hirashima and Dr N. Sato for the
generous gift of recombinant human G-CSF, produced by Chugai
Pharmaceutical Co (Tokyo, Japan). We also thank Dr Liv Theodorsen, MD
(The Central Laboratory, The Norwegian Radium Hospital, Oslo, Norway)
for letting us examine mouse blood with the blood cell analyzers.
| |
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Abstract
Introduction
Abstract