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Departments of Clinical Chemistry (M.V., E.D., A.N.M.) and Pharmacology (A.G.), University of Crete School of Medicine, Crete, Greece; and the Division of Endocrinology, Diabetes, and Metabolism, Department of Internal Medicine, University of Kentucky College of Medicine (K.A.), Lexington, Kentucky 40536-0084
Address all correspondence and requests for reprints to: Dr. Andrew N. Margioris, Department of Clinical Chemistry, University of Crete School of Medicine, Heraklion GR-711 10, Crete, Greece. E-mail: andym{at}med.uch.g
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Abstract |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Normal rat adrenal chromaffin and PC12 cells produce CRH (4). Our patient had ectopic Cushing’s syndrome with normal ACTH levels. Because of these data, we examined whether KAT45 cells produce CRH. In the second part of this report we describe our findings regarding the presence of the CRH transcript and immunoreactive (IR) CRH in KAT45 cells and their media. The transcript was characterized by Northern blot analysis, while the IR-material by sieve chromatography and RP-HPLC. The release rate of CRH from KAT45 cells was also assessed after exposure to nicotine and depolarizing concentrations of K+. The synthesis rate of CRH was examined after exposure to 7S NGF for 1 week, as it is now known that NGF, in addition to its effect on cell phenotype, also induces the expression of the CRH gene in primary pheochromocytoma cell cultures and in the PC12 cell line (4, 5). The effect of dexamethasone on CRH production was also tested, as glucocorticoids suppress the expression of the CRH gene in the hypothalamus but induce it in placenta (6). Finally, the effect of interleukin-1 (IL-1) was tested because this cytokine now ranks as a major stimulator of CRH production in the hypothalamus (7, 8) and the periphery. Furthermore, it should be noted that the IL-1 cytokines have been shown to be produced locally within the adrenal medulla and have been found to affect both normal adrenal chromaffin and pheochromocytoma cells (9, 10, 11, 12, 13).
Adrenomedullary CRH, in addition to its potential systemic effects,
which in our patient caused ectopic Cushing’s syndrome, may also exert
local paracrine-autocrine effects (4, 5). In the last part of this
report we present our findings regarding the paracrine effects of CRH
on KAT45 cells. Synthetic CRH and/or its antagonist 
-helical (h)
CRH were tested on the following parameters: 1) proliferation rate, as
CRH has been shown to affect the proliferation of normal anterior
pituitary corticotrophs (14) as well as that of the mouse corticotropic
cell line AtT20 (15); 2) production of catecholamines, as CRH has been
shown to directly stimulate the secretion of norepinephrine from
neurons in the prefrontal cortex (16, 17, 18, 19), hippocampus (20), and
hypothalamus (16, 17) and adrenaline from normal bovine adrenal
chromaffin cells (21); and 3) production of ACTH and ß-endorphin (4, 22), as CRH is a major regulator of POMC in pituitary, central and
peripheral nervous system, and several nonneuronal tissues that produce
it.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Sterile, fresh tissue, from within the tumor capsule was minced and cultured in flasks containing antibiotic-free RPMI 1640 medium with 10% (vol/vol) FBS (all from Life Technologies, Gaithersburg, MD) in humidified 5% CO2 at 37 C.
PC12 cells
The PC12 cells were obtained from two sources: 1) Dr. G. Guroff,
Section on Growth Factors, NICHHD, NIH (Bethesda, MD); and 2) the
American Type Culture Collection (Rockville, MD). The conditions of
their handling in our laboratory have been previously described (2, 3, 4).
The PC12 cells were cultured on either flat bottom wells in 12-well
plates of 4.5-cm2 surface area/well (Costar Europe,
Badhoevedorp, The Netherlands) or on flasks of 75-cm2
surface area, at an initial concentration of 1 x 105
cells/cm2. They were left to grow in Eagle’s MEM (Life
Technologies) containing 10 mM L-glutamine, 1.5
mM HEPES, 100 U/ml penicillin, 0.1 mg/ml streptomycin,
7.5% horse serum, and 7.5% FCS in an incubator (Forma Scientific,
Marietta, OH) at 5% CO2 and 37 C. Differentiation of PC12
to neuron-like cells followed exposure to 7S NGF (25 ng/ml x 1 week).
The 7S NGF was provided by Dr. G. Guroff, Section on Growth Factors,
NICHHD, NIH.
Proliferation
The proliferation rate was determined under basal conditions and
after exposure to CRH. It was measured by 1) cell counting, 2) the MTT
(Sigma Chemical Co., St. Louis, MO) method, and 3)
[3H]thymidine incorporation. For cell counting, cells
were trypsinized, centrifuged, resuspended in RPMI and 10% FCS, and
counted visually on a hematocytometer using trypan blue. In addition,
the cells were counted on a desktop Coulter counter (Coulter
Electronics, Hialeah, FL). For the MTT method, the cells were plated in
flat bottom 96-well plates at an initial concentration of 6,000-10,000
cells/well on RPMI-10% FBS. MTT was dissolved in PBS at 5 mg/ml (stock
solution). KAT45 cells were plated in flat bottom wells (96
wells/plate) at an initial concentration of 6,000–10.000 cells/well,
on RPMI medium containing 10% FBS. The next day (second day of the
experiment), this medium was replaced by test medium free of serum
containing CRH, hCRH, or their vehicles (controls). At the end of
each experiment, the media were replaced by fresh test medium. The
cells were left to rest for 1 h, and MTT was added at a final
concentration of 0.5 mg/ml and incubated for an additional 4 h at
37 C. Living mitochondria metabolize MTT to blue crystals. The MTT
media were then replaced by a solution of acid-isopropanol, and the
optical density was read on a Dynatech MicroElisa reader (Chantilly,
VA) at a wavelength of 600 nm. For the incorporation experiments, the
cells were aspirated from 6-well plates and transferred to flat bottom
96-well microtiter plates (Costar, Cambridge, MA). Each well was then
pulsed with 1 µCi [methyl-3H]thymidine (SA,
25 Ci/mmol; Amersham Life Science, Aylesbury, UK) for 24 h (a 24-h
incubation was found to give the best incorporation) at 37 C in 5%
CO2-95% air (3). At the end of the final incubation, the
cells were aspirated automatically by a cellular harvester (Minimash
2000, Dynatech, France), transferred to prewet filters, vacuum-washed
three times, dried for 90 min at 65 C, cut, and counted in a liquid
scintillation ß-counter (4000 series, United Technologies, Packard,
Downers Grove, IL).
Separation and measurement of catecholamines
The separation and measurement of catecholamines in culture
medium were performed as previously described (2). Briefly, samples of
600 µl were aspirated from each well and transferred to tubes
containing 100 µl 0.1 N HCl to ensure the stability of
the catecholamines (stable in acidic pH). Subsequently, the
catecholamines were extracted by acetone precipitation. The extracts
were then evaporated and reconstituted in 200 µl of the mobile phase
of the HPLC. Samples of 25 µl (i.e. one eighth of the
initially extracted material) were injected into the RP-HPLC (HP-1090,
Hewlett-Packard, Waldbronn, Germany) equipped with an electrochemical
S100A detector (ESA, Coulochem, Bedford, MA). The detector was set at
+0.45 mV (with respect to an H2/H+ couple
reference electrode), and the guard cell was set at +0.5 mV. The guard
and main columns were RP-18, 5-µm particle size (Hewlett-Packard),
whereas the size of the main column was 200 x 4.6 mm. The mobile
phase was composed of 25 mM
NaH2PO4·H2O, 3.2 mM
1-heptane sulfonic acid sodium salt, 0.5 mM EDTA, and 7.5%
methanol. The signals were recorded in an analog digital converter
(HP-35900 C, Hewlett-Packard) connected to an HP-1040 computer.
CRH, ACTH, and ß-endorphin assays
Peptides in culture medium were concentrated by C18
reverse phase columns (Sep-Pak, Waters Associates, Milford, MA) after
acidification in 2 vol 0.1 N HCl and centrifugation at
1.5 x 103 rpm for 10 min. The supernatants were
extracted by activated Sep-Pak cartridges, washed with 20 ml 0.1
N HCl, eluted with 3-ml acetonitrile 0.01–80% HCl, then
dried under vacuum (Speed-Vac, Savant Instruments, Farmingdale, NY).
The CRH content of the Sep-Pak extracts was assayed by RIA using a
rabbit antiserum purchased from Neosystem Laboratoire (Strasbourg,
France). The antiserum was raised against human CRH and exhibits 100%
cross-reactivity to rat CRH and no cross-reactivity to ovine CRH, human
ACTH, human ß-endorphin, LHRH, or arginine vasopressin. The
sensitivity of the assay was 1 pg/tube. The intraassay coefficient of
variation was 4.4%, and the interassay coefficient of variation was
6.6%. Results are expressed as picograms of IR-CRH per mg total
cellular protein determined on whole cellular homogenates. ACTH in the
culture medium was measured by chemiluminescence (Nichols Institute
Diagnostics, San Juan Capistrano, CA). An acridinium ester-labeled
mouse monoclonal ACTH-(1–17) antibody and a biotin-coupled goat
polyclonal ACTH antibody exhibiting 100% cross-reactivity to
ACTH-(1–39) and no cross-reactivity to ACTH-(1–24), ACTH-(1–10),
MSH, and ß-endorphin were used. The sensitivity of the assay was
0.5 pg/ml medium. The intraassay coefficient of variation was 3.4%,
and the interassay coefficient of variation was 4.6%. Results are
expressed as picograms of ACTH per mg total cellular protein. For the
measurement of IR-ß-endorphin, a previously described protocol was
used (23). Briefly, culture media were first acidified in 5 vol 0.1
N HCl and centrifuged at 10,000 rpm for 20 min, and the
supernatant was passed through Sep-Pak cartridges (Waters Associates,
Milford, MA), eluted by 0.01 N HCl-80% acetonitrile in
tubes containing 1 mg human serum albumin, and then dried by Speed-Vac
and reconstituted in RIA buffer. The assay was a solid phase two-site
immunoradiometric assay (Nichols Laboratories, Irvine, CA). The
antiserum was raised again human ß-endorphin and exhibits a 16%
cross-reactivity to ß-lipotropin and no cross-reactivity to ACTH,
MSH, - or ß-neo-endorphin, or the enkephalins and dynorphins.
Under our conditions, half-maximal tracer displacement occurred between
20–25 pg, and the sensitivity was 1–2 pg/tube. Serial dilution of a
pool of the extracted culture media displaced the radioactive tracer
from antiserum in parallel with the synthetic ß-endorphin.
Sieve chromatography and RP-HPLC
The mol wt of IR-CRH was assessed by gel filtration
chromatography. Briefly, pooled culture media were acidified in 2 vol
0.1 N HCl and centrifuged at 1 x 105 rpm
for 10 min, and the supernatant was extracted by activated Sep-Pak
cartridges, washed with 20 ml 0.1 N HCl, eluted with 3 ml
acetonitrile 0.01–80% HCl, and dried under vacuum (Speed-Vac). The
samples were reconstituted in 0.5 ml 10% formic acid containing 0.5%
defatted BSA and 6 M urea and chromatographed on Sephadex
G-50 (0.9 x 60 cm; bed volume, 40 ml; flow rate, 1.5 ml/h; 1-ml
fractions collected). Fractions were dried under vacuum and
reconstituted for RIA. After chromatographies of the samples, the G-50
column was calibrated with blue dextran and synthetic human CRH.
For the characterization of IR-CRH by RP-HPLC, KAT-45 cells in culture
were scrapped, pooled, homogenized in 5 vol 0.1% trifluoroacetic acid
(TFA), and centrifuged at 10,000 rpm for 20 min; the supernatant was
dried under vacuum, were reconstituted in 30 µl 25%
acetonitrile-0.1% TFA, and centrifuged again for 10 min; and one third
of the supernatant was injected into the HPLC connected to a 0.2
x 30-cm C18 µBondapack column (Waters Associates,
Milford, MA). A linear gradient program was used employing the
TFA-acetonitrile solvent system shown in Fig. 3
. Solvent A was composed
of 20% acetonitrile-0.1% TFA, and solvent B was composed of 80%
acetonitrile-0.1% TFA. The flow rate was 1 ml/min. Samples of 1 ml
were collected. The retention time (Rt) of the IR-CRH
extracted from the KAT45 cells was determined by RIA after evaporation
of the HPLC fractions and reconstitution in RIA medium. After the
sample runs, 20 ng diluted synthetic CRH and CRH oxidized by 5%
H2O2 were run, and their Rt values
were determined by RIA.
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-32P]deoxy-ATP (1200 Ci/mmol; Amersham,
Arlington Heights, IL) and terminal deoxynucleotidyl transferase
(Boehringer Mannheim) to a specific activity of 0.8 x
10-9 dpm/mg. Hybridization was performed in the
prehybridization buffer containing 1 x 10-6 dpm/ml
labeled CRH probe at 42 C. Blots were washed in 0.2 x SSC-0.1%
SDS for 1 h at 42 C and exposed at -70 C to Kodak XR film
(Eastman Kodak, Rochester, NY) in the presence of intensifying
screens.
Statistical analysis
The concentration of IR-CRH in the culture medium was normalized
according to the total cellular protein content after resuspension and
homogenization at the end of each experiment (4). The secretion of
epinephrine and dopamine in the culture medium is expressed as either
the mean (nanograms) ± SEM of five or six experiments (per
well per mg total cell protein) or as percent changes compared with
control wells. Total cellular protein in each well was measured by a
modification of the Bradford Coomassie Brilliant Blue G250 method
(Serva, Heidelberg, Germany) using BSA as standard (25). For the
statistical evaluation of our data, we used ANOVA followed by multiple
comparison Fisher’s least significance difference test and the
Newman-Keuls test. For data expressed as the percent change over
controls, we used the nonparametric test of Kruskal-Wallis for several
independent samples.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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depicts KAT45 cells
24 h after plating at low power. KAT45 cells exhibit a size
variation that tends to disappear as cells grow. Figure 1B depicts a
high power view of KAT45 cells, the phenotype of which remained
unchanged after freezing and thawing. Figure 1C depicts a high power
view of PC12 cells; the morphological features and size of these cells
resemble those of KAT45 cells. Figure 1D depicts the effect of 7S NGF
at 25 ng/ml for 1 week on KAT45 cells, whereas Fig. 1E shows its effect
on PC12 cells. Figure 2, depicts the
proliferation rate of KAT45 cells under basal conditions measured by
cell counting of trypan-excluding cells (Fig. 2A). The doubling time,
under our culture conditions, was approximately 24–48 h, which is
similar to that of PC12 cells (data not shown). MTT assays of the
steepest part of the proliferation curve confirmed these results (Fig. 2B, upper curve).
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depicts the effect of nicotine at 10-6 M; it
caused a highly significant increase in norepinephrine release at 15
min that lasted up to 60 min.
Production of CRH
Figure 4 depicts the concentration
pattern of IR-CRH in the medium of KAT45 cells over a period of 3 days
under basal conditions. It also shows the characterization of the
released IR-peptide and its transcript. Specifically, Fig. 4A depicts
the concentration of IR-CRH in the culture medium 3, 6, 9, 12, 12, 48,
and 72 h after the removal of FBS-enriched culture medium and its
replacement by plain medium containing aprotinin and BSA. The
concentration of IR-CRH in the medium exhibited a wave-like pattern, a
phenomenon also observed in other chromaffin cells (4). The mean
concentration of IR-CRH ranged from 20–30 pg/mg total cellular protein
or approximately 0.5–1 x 10-8 M. The
inset of Fig. 4A depicts the elution profile of
KAT45-derived CRH on sieve chromatography, which is similar or
identical to that of synthetic human CRH. Figure 4B depicts the RP-HPLC
chromatographic profile of KAT45-derived IR-CRH. A gradient of
acetonitrile (CH3CN) was used (indicated by the
broken line) in the presence of 0.1 N TNF. The
IR-CRH from the KAT45 cells had two peaks. The first had the
Rt of oxidized Met(O) CRH (formed during the extraction
procedures), whereas the second peak had the Rt of
nonoxidized human/rat CRH standard. The inset of Fig. 4B (at
the top) depicts the Northern blot signal of KAT45 total RNA
after hybridization with a 42-mer CRH oligo (right) compared
with the signal of total RNA from human placenta (left). The
calculated size of both hybridization signals was the same as that of
the authentic transcript.
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depicts the
effect of nicotine (10-6 M) on CRH compared
with levels in parallel controls. The stimulatory effect of nicotine on
CRH peaked at 40 min, whereas its effect on norepinephrine peaked
earlier, at 15 min. Although the costorage and corelease of classical
neurotransmitters with neuropeptides is a characteristic of neural
crest-derived cells, the concentration of neurotransmitters in the
culture medium depends not only on the rate of their release but also
on the rate of their reuptake (26, 27). Thus, the asynchronous peaks we
find may be due to reuptake of norepinephrine by KAT45 cells. Figure 5B
depicts the effect of K+-induced depolarization, which
peaked at 20 min.
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).
Similarly, IL-1ß at 10 ng/ml exerted a highly significant stimulatory
effect only after 24 h or more (Fig. 6B). Nerve growth factor
(NGF) also appeared to involve CRH synthesis. Figure 6C depicts the
effect of 7S NGF (25 ng/ml x 1 week) on CRH production compared with
that in parallel controls (KAT45 cells cultured in parallel for the
same number of days and not exposed to NGF). PC12 rat and primary
cultures of human pheochromocytoma cells respond to NGF in a similar
manner (4, 5).
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, lower curve, depicts the suppressive effect of CRH at
10-6 M on the proliferation of KAT45 cells.
The trend of declining proliferation became statistically significant
by the fourth day of culture. This effect of CRH appears to be
pharmacological rather than physiological, as lower concentrations of
CRH were ineffective (data not shown). Furthermore, the specific
antagonist hCRH did not have any effect of its own (data not shown),
suggesting that endogenous CRH does not play a role in the
proliferation process of KAT45 cells.
CRH stimulates the synthesis of catecholamines in the locus coeruleus
(19), hippocampus (20), prefrontal cortex (16, 18), hypothalamus (16, 17), adrenal chromaffin (4, 21), and PC12 rat pheochromocytoma cells
(4). CRH had a similar stimulatory effect on KAT45 cells. A short
exposure of KAT45 cells to CRH (<12 h) did not cause any significant
effect, suggesting that CRH does not influence the release of
catecholamines, a finding in agreement with previously published data
(4, 21). Longer exposure (36 h or more) resulted in a highly
significant stimulation, suggesting the involvement of biosynthetic
mechanisms. Figure 7A depicts the
dose-dependent stimulatory effect of CRH on norepinephrine secretion
from KAT45 cells exposed to it for 36 h (upper curve).
The simultaneous addition of the CRH antagonist hCRH in excess
(10-5 M) abolished the stimulatory effect of
CRH to a significant degree (lower curve), suggesting the
involvement of specific CRH receptors. However, hCRH did not have
any significant effect of its own (data not shown).
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shows
that the stimulatory effect of CRH on IR-ß-endorphin release from
KAT45 cells was dose dependent, whereas Fig. 7C shows that its
time-dependent response peaked at 9 h. Similar responses were
obtained for ACTH (data not shown). |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Our patient had ectopic Cushing’s syndrome with normal ACTH levels, a finding suggesting that its cause was ectopic production of CRH from the tumor and release into the systemic circulation. However, it is now widely suspected that the production of CRH by pheochromocytomas may be a rather common phenomenon. On the other hand, the development of Cushing’s syndrome from the systemic action of tumoral CRH is probably quite rare. Although we know of no study that evaluates the percentage of pheochromocytomas producing pathophsiologically relevant amounts of CRH, it appears that in most cases CRH produced by this type of tumor does not exert any systemic effect. Indeed, CRH has been shown to be produced in primary cultures of human pheochromocytomas, a small minority of which caused ectopic Cushing’s syndrome (28, 29, 30, 31, 32, 33). Furthermore, it now appears that CRH is also produced by normal human adrenal medulla (28, 29, 34, 35) and by the medulla of rodents (11, 36, 37, 38), cows (39, 40, 41), and dogs (42). Based on these data we have hypothesized that the production of CRH may represent the preservation of a physiological characteristic of adrenal chromaffin cells rather than a tumor-induced phenomenon. Indeed, dispersed normal rat adrenal chromaffin cells and the PC12 rat pheochromocytoma cell line produce comparable amounts of CRH (4). In the few cases in which there is development of Cushing’s syndrome from a systemic effect of CRH, we beleive that this reflects a failure of the normal barriers against it rather than a dramatic elevation of its concentration in plasma. The precise mechanism of this failure is not readily apparent, but it may be an enhanced bioavailability of CRH secondary to a decreased concentration of the CRH-binding proteins or a parallel increase in the sensitivity of anterior pituitary corticotrophs to CRH.
It is now well established that the immune system cross-talk with the two stress axes, the hypothalamic-pituitary-adrenal axis and the sympathetic system (central and peripheral). Indeed, systemic and local macrophage-derived IL-1 cytokines affect the two axes centrally at the level of the hypothalamus (where they stimulate the production of CRH) and at the level of the locus coeruleus (where they stimulate the production of noradrenaline) (7, 8, 43). In the periphery, exposure of normal adrenal medulla to recombinant human IL-1ß increases both the production of CRH and that of catecholamines (4, 12). Our findings are in complete agreement with these data. Our human pheochromocytoma cells respond to recombinant human IL-1ß by increasing the rate of their CRH synthesis, suggesting that the response of normal adrenal chromaffin cells to these cytokines may be preserved in some pheochromocytomas.
It is now known that in a growing number of extracranial sites, glucocorticoids stimulate the expression of the CRH gene instead of suppressing it. For example, it has been found that in primary cultures of human placental cytotrophoblasts (6) and in human pheochromocytomas (5), glucocorticoids stimulate the production of CRH. In agreement with these data, we have found that glucocorticoids stimulate the production of CRH in our cells. The physiological significance of this phenomenon and the mechanism involved are currently speculative.
Several reports have shown that NGF affects the production of CRH from neural crest-derived cells. Indeed, NGF increases the CRH messenger RNA content of primary cultures of human pheochromocytomas by 10-fold (5) and the production of the CRH peptide by PC12 rat pheochromocytoma cells (4). This phenomenon may be associated with the acquisition of neuron-like characteristics after exposure to NGF (4). Our KAT45 data strengthen these observations, suggesting that the NGF-mediated neuronal transformation of pheochromocytoma cells may be functionally linked to the expression of the CRH gene, as the two phenomena appear to develop in parallel. It is possible that CRH may mediate some of the effects of NGF. However, preliminary data on the effect of CRH or its antagonist on the change in our cell phenotype were not promising. Thus, it is possible that the changes in CRH production observed after NGF treatment might be a nonspecific phenomenon, part of a major reorganization of the cellular metabolism initiated by NGF. Indeed, NGF-treated cells look completely different from parallel controls.
It is our hypothesis that the biological role of peripherally produced
CRH is local, and any potential systemic effect must first overcome the
protective barrier of the circulating CRH-binding protein. A local
effect of CRH on adrenals and their tumors is probable because they
both contain specific CRH-binding sites (21, 44, 45). In agreement with
this hypothesis, we have found that several parameters of PC12 and
KAT45 cells appear to be affected by CRH, including their proliferation
rate (a rather pharmacological effect), production of catecholamines,
and POMC-derived peptides. In our hands, the stimulatory effect of CRH
on the production of catecholamines was dose and time dependent and was
blocked by the specific antagonist hCRH, suggesting the mediation of
CRH receptors. In this regard, the response of our cells to CRH was
similar to that of normal chromaffin cells (21) and catecholaminergic
neurons in the central nervous system (16, 17, 19, 20). In general, CRH
is a potent effector of catecholamine metabolism, stimulating the
synthesis of tyrosine hydroxylase (46, 47), the rate-limiting enzyme in
their synthetic pathway. In the central nervous system, the stimulatory
effect of CRH on the production of catecholamines appears to be exerted
in a paracrine manner via the close anatomical proximity of the CRH and
the noradrenergic neurons (48, 49). Thus, the close association
between the hypothalamic-pituitary-adrenal and catecholaminergic
stress axes appears to be retained in both the normal adrenal
medulla and at least some pheochromocytomas.
CRH affects the proliferation of normal anterior pituitary
corticotrophs (14) and the corticotropic cell line AtT20 (15). We found
that CRH suppressed the proliferation of KAT45 cells. However, this
effect does not appear to be of physiological significance, as it
occurred only at the highest dose of CRH, and hCRH, the specific CRH
antagonist, did not have any effect of its own, suggesting that KAT45
cells were not under tonic inhibition by endogenous CRH as in the case
of endogenous opioids in which the addition of opioid antagonists
exerts a major effect on their proliferation (3).
In conclusion, our KAT45 cells, in addition to the data they provided regarding the in vitro profile of a human CRH-producing pheochromocytoma, may prove to be a valuable auxiliary to the PC12 cell line in the study of neuronal differentiation and the neuro-immuno-hormonal interactions regulating the production of catecholamines in normal and tumorous adrenal medulla.
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Footnotes |
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2 Present address: Division of Endocrinology, Department of
Pediatrics, Childrens Hospital, Harvard Medical School, 300 Longwood
Avenue, Boston, Massachusetts 02115.
Received April 11, 1997.
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