Published in Volume
119, Issue 5
(May 1, 2009)J Clin Invest.
Copyright © 2009, American Society for Clinical
Substance P stimulates human airway submucosal gland secretion mainly via
a CFTR-dependent process
1Cystic Fibrosis Research Laboratory, Stanford University, Stanford,
2Department of Otorhinolaryngology, Yonsei University,
Seoul, Republic of Korea.
3Department of Pediatrics, UCSF, San Francisco,
4Department of Cardiothoracic Surgery and
5Department of Pulmonary and Critical Care Medicine, Stanford University
School of Medicine, Stanford, California, USA.
Address correspondence to: Jeffrey J. Wine, Cystic Fibrosis Research
Laboratory, Room 450, Bldg. 420, Main Quad, Stanford University, Stanford, California
94305-2130, USA. Phone: (650) 725-2462; Fax: (650) 725-5699; E-mail:
First published April 20, 2009
Received for publication August 27,
2008, and accepted in revised form February 25,
Chronic bacterial airway infections are the major cause of mortality in cystic
fibrosis (CF). Normal airway defenses include reflex stimulation of submucosal gland
mucus secretion by sensory neurons that release substance P (SubP). CFTR is an anion
channel involved in fluid secretion and mutated in CF; the role of CFTR in secretions
stimulated by SubP is unknown. We used optical methods to measure SubP-mediated
secretion from human submucosal glands in lung transplant tissue. Glands from control
but not CF subjects responded to mucosal chili oil. Similarly, serosal SubP
stimulated secretion in more than 60% of control glands but only 4% of CF glands.
Secretion triggered by SubP was synergistic with vasoactive intestinal peptide and/or
forskolin but not with carbachol; synergy was absent in CF glands. Pig glands
demonstrated a nearly 10-fold greater response to SubP. In 10 of 11 control glands
isolated by fine dissection, SubP caused cell volume loss, lumen expansion, and mucus
flow, but in 3 of 4 CF glands, it induced lumen narrowing. Thus, in CF, the reduced
ability of mucosal irritants to stimulate airway gland secretion via SubP may be
another factor that predisposes the airways to infections.
Cystic fibrosis (CF) is caused by mutations that disrupt CFTR, an anion channel that
mediates fluid secretion in many epithelia. The most serious consequence of CF is
respiratory failure caused by chronic bacterial airway infections, which motivates
research to understand the link between loss of CFTR function and airway infections. In
the airways, CFTR is found in both the surface ciliated epithelial cells and in serous
cells of submucosal glands (1–3). At both sites, it helps mediate fluid secretion
that hydrates mucins and other innate defense molecules elaborated primarily by the
glands (4, 5). The resulting bacteriostatic mucus is efficiently transported out of the
airways, keeping normal airways sterile. In CF, CFTR-dependent anion and fluid secretion
by both surface epithelia and glands is lost (6,
7), but absorption is increased (reviewed in
ref. 8), and it has been hypothesized that
hyposecretion/hyperabsorption leads to defective mucus clearance (9) that compromises innate mucosal defenses and predisposes CF
airways to chronic infections (5, 7, 10–24). Fortunately,
CFTR-independent mechanisms exist for fluid secretion in both surface epithelia and
glands so that some innate defense persists. Current treatments for CF airways disease
often engage these CFTR-independent secretion pathways (8), and such strategies might be improved if we had a better understanding of
CFTR’s role in airway innate defense.
Submucosal glands produce most upper airway mucus (25, 26), and in experiments using
tracheal xenografts, with and without glands, the gland-containing xenografts elaborated
much more lysozyme and were much more effective in resisting infections (27). Airway glands are controlled by autonomic
efferent innervation from airway intrinsic neurons that release multiple
neurotransmitters, including acetylcholine (ACh), vasoactive intestinal peptide (VIP),
substance P (SubP), and NO, as well as by mediators like histamine, bradykinin, and
PGE2 (reviewed in ref. 17). Glands
also receive SubP via axon reflexes from C fiber afferents. We are attempting to
understand the roles these different agonists play in controlling the glands and the
extent to which the secretion they stimulate is CFTR dependent; such information is
essential if we are to understand the role of CFTR-mediated secretion in airway innate
In prior studies, we found that while ACh can still stimulate secretion from CF airway
glands, fluid secretion stimulated by VIP is absent (7). The difference was attributed to the ability of ACh to elevate intercellular
Ca2+ concentrations ([Ca2+]i) and engage non-CFTR
anion channels, and this has now been validated in studies of mouse submucosal glands
(28, 29). SubP, like ACh, appears to elevate [Ca2+]i in most
systems, and in pig bronchi, SubP is more potent and has similar efficacy as ACh in
stimulating gland mucus secretion (26, 30). This suggests that SubP, like ACh, may act in
part via a Ca2+-dependent, CFTR-independent process in pigs. However, when
testing mucus secretion from airway glands in mice, Ianowski et al. found that SubP
stimulation was a much less effective agonist than cholinergic stimulation (31) and that unlike cholinergic stimulation, which
produces equivalent secretion in WT and
Cftr–/– mice (18), SubP was completely ineffective in producing
mucus secretion in Cftr–/– mice
(31). Hyposecretion of mucus by CF airway
glands may seem counterintuitive, given the copious sputum that is a hallmark of CF
airways disease. However, there is a growing realization that excess sputum occurs in
part as a consequence of overproduction secondary to chronic airway infection and
inflammation and in part due to increased difficulty of clearing the CF sputum
unobtrusively via mucociliary clearance and swallowing (32).
Given these large species differences in airway gland responses of mice and pigs to
SubP, it became essential to determine the responses of human airway glands to SubP.
Surprisingly, in the present work, we show that humans resemble mice more than pigs,
with regard to their airway gland responses to SubP. In humans as in mice, SubP is a
much less effective agonist than ACh (carbachol), and overt fluid secretion stimulated
by SubP is almost completely lacking in human CF glands, although morphological changes
in the CF glands can still be observed using differential interference contrast (DIC)
optics. Based on synergy between SubP and saturating concentrations of forskolin, we
suggest a model in which CFTR-dependent secretion stimulated by SubP is produced by a
mechanism other than merely elevating [cAMP]i.
Human control glands but not CF glands secrete in response to mucosal chili oil. When droplets of chili oil were added to the mineral oil layer covering the mucosal
surface of isolated human airway segments, quiescent glands from human normal (HN) or
disease control (DC) tissues began to secrete, but glands from CF airways did not
(Figure 1). This result in humans recapitulates
previously reported results for control and
Cftr–/– mice (18). In any preparation, only a portion of viable
glands responded (viability was tested with carbachol applied to the serosal side),
and the average secretion rates for responding glands were approximately 10% of rates
of response to 10 μM carbachol.
In human airways, mucosal chili oil stimulates submucosal gland mucus
secretion from control (HN and DC) but not CF subjects. (A) Images of mucus bubbles formed under oil at the orifices of
single submucosal glands in response to stimulation via mucosal chili oil. At 15
minutes, 2 μl of chili oil was added to the approximately
20–30 μl oil layer. No mucus secretion was seen during the
15-minute control period (left panel); the right panel shows the results 20
minutes later. Scale bar: 0.5 mm. (B) Summary data showing average
secretion rates (± SD) for 20-minute periods, following application of
chili oil. If basal secretion was present in the control period, it was subtracted
from the rates shown. The secretion rate for CF glands was less than control
glands (*P < 0.05 versus HN + DC).
Human control glands responded to SubP. SubP is the most likely transmitter for mediating the secretory responses to mucosal
chili oil. Therefore, we applied SubP directly to the serosal side of our human
airway preparations, where it was able to access the glands directly. The results of
these experiments were consistent with the results using chili oil. When isolated
segments of tracheal or bronchial airways were stimulated with 1–100
μM of serosal SubP, single glands started to secrete within
1–2 minutes and some but not all glands continued to secrete as long as
SubP was present (Figure 2A). Only a portion of
viable glands responded (Figure 2C). For 24 HN
subjects, 100% of subjects and 295 of 379 of viable glands responded. The proportion
of viable glands responding to SubP per HN subject varied between
33%–100% (mean, 78% ± 16%). For 20 DC subjects, 95% of
subjects and 175 of 293 of viable glands responded (25%–100% per
responding subject; mean, 60% ± 18%).
Serosal application of SubP-stimulated secretion from HN and DC but not CF
airway glands. (A and B) Plots of secreted mucus volume over time for
individual glands from 1 DC and 1 CF subject after stimulation are shown (each
line represents a single gland). Right panels show part of the fields at end of 20
minutes of basal, SubP, and carbachol (carb) stimulation, respectively. Scale bar:
0.5 mm. (C) SubP-stimulated secretion in a larger proportion of
glands in HN or DC versus CF subjects. Each bar shows the mean percentage of
viable glands that responded to SubP in HN, DC, and CF subjects. The total number
of viable glands was defined as the number responding to 5 minutes of 10
μM carbachol at the end of the experiment. Error bars show SEM. CF
response to SubP was significantly less than that of either HN or DC;
*P < 0.05.
A concentration-response relationship was established by measuring the amount of
secretion during the first 15 minutes of stimulation with SubP in the constant
presence of 1 μM of phosphoramidon (Figure 3A). For this method, Vmax = 0.33
± 0.02 nl/min/gland and EC50 = 1.13 ± 0.22
μM. Maximal secretion rates for responding glands in controls were
approximately 10% of those seen with 10 μM carbachol (Figure 3, B and C, and Figure 4A).
Secretion rates of human airway glands to SubP. (A) Approximate dose-response relationship for submucosal gland
secretion from HN glands. Each point is the average of 10–14 glands
from 2–3 different HN subjects; 5 HN subjects were used for the graph,
and all data were obtained within 8 hours of harvest. The secretion rate was
averaged over the first 15 minutes following SubP addition. Up to 3 increasing
concentrations were administered per subject. (B and C)
Summary data for average secretion rates measured 15 minutes after 10
μM SubP and 5–10 minutes after 10 μM
carbachol. The scale for responses to carbachol is 10 times the scale for
responses to SubP. Bars in B show the secretion rate for responding
glands and are labeled with the number of subjects, number of responding glands,
and total number of glands tested. Five CF glands responded to SubP of more than
100 tested. The CF secretory rate for the 5 responding glands was significantly
less than that for either HN or DC glands. *P < 0.05. Bars
in C show the secretion rate for responding glands and are labeled
with the number of subjects and total number of glands tested. Error bars are SEM
Typical gland response to SubP. (A) The time course of secretion rate to SubP shows an early peak
like the response to carbachol. The graph shows (mean ± SEM) secretion
rates averaged over 5-minute periods for 15 glands from 3 HN subjects tested
6–9 hours from harvest; 10 μM SubP plus 1 μM
phosphoramidon was added at 10 minutes. The inset is rescaled to show, in these
same glands, both the response to SubP (open arrow) and the subsequent response to
10 μM carbachol, added at the 45-minute time point (arrow).
(B) Age- and disease-related changes in responses of human airway
glands to SubP in HN, DC, and CF subjects. Each symbol shows the percentage of
glands that responded to SubP as a function of type and age of subject. Data are
for a subset of subjects, in which counts were made of total glands responding to
The time course of secretion stimulated by SubP characteristically had an early peak,
followed by a more sustained response that was approximately one-third or less of the
peak secretory rate (Figure 4A). This response
profile essentially duplicated that seen for carbachol in sheep (33), pig (34) and human glands (see below), although the magnitude of the response was
Human CF glands barely responded to serosal SubP. In contrast with controls, a significantly smaller proportion of viable glands in CF
subjects responded to SubP (Figure 2, B and C,
and Figure 3B). The mean percentage of
responding CF glands was only 4% ± 4%, about 5% of the control
percentages (n = 7; only 5 of 112 viable glands responded; range,
0%–9% per subject with 4 subjects not responding). In the CF tissue with
the highest proportion of responding glands (CF48, ΔF508 homozygous), 2
of 22 glands (9%) secreted in response to SubP. For these 2 responding glands, the
peak secretion rates observed were 3% and 6% of the subsequent responses to
carbachol, both less than the 10% seen for control glands (Supplemental Figure 1;
supplemental material available online with this article; doi:10.1172/JCI37284DS1).
As will be shown below, a single gland acinus isolated from this same subject for DIC
imaging showed clear (but transient) acinar secretion.
In the above analyses, we averaged secretion rates only for responding glands. The
total volume of secretion for a unit area of membrane is equal to the number of
responding glands multiplied by the gland secretion rate. The values in Figures 2 and 3
indicate that the total volume of secretion stimulated by SubP for CF tissues is
approximately 0.4% of that control tissues.
Responses to SubP decreased with patient age in control subjects. Some evidence indicates possible age-related changes in airway innervation by
peptide-containing nerves (35), so we plotted
responsiveness as a function of patient age in HN, DC, and CF subjects. In controls,
the proportion of responding glands decreased with age and with the presence of other
diseases (Figure 4B). Because DC subjects were
older on average than HN subjects, with little overlap (Figure 4B and Table 1), we are
unable to determine conclusively whether age was the major variable accounting for
the slight difference between the 2 groups. However, the youngest DC subject had a
secretion rate equivalent to similarly aged HN subjects, and the oldest HN subject
had a secretion rate equivalent to similarly aged DC subjects. By contrast, the CF
subjects, who were on average the youngest, had by far the lowest proportion of
responding airway glands (Figure 4B).
Subject characteristics for individual gland secretion rate experiments
Rates of mucus secretion stimulated by SubP and carbachol were correlated. Because some control glands did not respond to SubP, we considered the possibility
that a distinct set of SubP-responsive glands exists. To test this idea, we plotted
peak secretion rates to SubP and carbachol on a gland-by-gland basis and obtained a
positive correlation between the 2 (r = 0.76, P
< 0.0001; Figure 5). The different axes
reflect the higher potency of carbachol, and the least squares line fit intercepted
the x axis for responses to SubP at a secretion rate of
approximately 2 nl/min/gland for carbachol. (For 18 glands with a measured response
of 0.0 to SubP, the mean secretion rate to carbachol was 1.3 ± 0.2
nl/min/gland.) The relationship plotted in Figure 5 suggested that no special set of SubP responsive glands exists but rather
that SubP is a weak agonist for all airway glands, so that glands with intrinsically
lower secretion rates do not produce measurable mucus at the gland duct orifice. This
result is consistent with prior results with carbachol and forskolin, in which we
noted a large range of secretion rates among glands to both agonists (7, 33),
with rates being positively correlated (34).
Secretion rate to SubP is positively correlated to secretion rate to
carbachol. Each point shows the peak single gland secretion rates to carbachol and SubP from
21 glands from 4 HN subjects and 13 glands from 3 DC subjects. One symbol is used
for both groups, because their rates did not differ. SubP stimulation preceded
carbachol. Of 110 glands, 18 (16%) responded only to carbachol; the mean peak
secretion rate of those glands was 1.26 ± 0.18 nl/min/gland. The solid
line shows linear regression (r = 0.76, P
Gland secretion stimulated by SubP was synergistic with VIP and/or forskolin for
control glands but not for CF glands. We observed synergistic increases in secretion rates when SubP was added to a
saturating dose (10 μM) of forskolin in HN or DC control glands (Figure
6, A and C). In contrast, CF glands showed
almost no measurable secretion stimulated by these agents alone or in combination
(Figure 6, B and C). In Figure 6C, we normalized all secretion rates to peak
secretion rates of carbachol; thus, the secretion rates are shown as fractions of the
carbachol responses. Normalization decreases the variability inherent in the
secretory responses of individual glands (see individual rates in Figure 6, A and B) and is appropriate because all groups
have similar responses to carbachol (see Figure 3C). The addition of SubP to VIP and/or forskolin also increased the
proportion of responding glands in 7 of 8 control subjects but not in any of 3 CF
subjects tested in this way (Supplemental Figure 2).
Stimulation of secretion by SubP is synergistic with forskolin or VIP in HN
and DC but not CF glands. (A) Secretion from individual HN glands to 10 μM
forskolin (forsk), 10 μM SubP plus forskolin, and finally 10
μM carbachol. (B) The same sequence in a CF subject
except for a 1-hour interval of basal secretion (data not shown). (C)
Summary showing synergism in control subjects for saturating concentrations of
SubP (averaged over 15 minutes), with saturating concentrations of either
forskolin or VIP (averaged over 20–30 minutes). Responses were
normalized to the peak secretory rates of those same glands to 10 μM
carbachol given at the end of the experiment. Responses to forskolin plus SubP in
HN (white bars) or DC (gray bars) subjects were greater than the summed responses
of the agents given separately (indicated by the dashed lines; *P
< 0.05). In contrast, CF glands (black bars) showed minimal responses to
either agent alone or in combination; (**P < 0.01 versus
control groups). Experiments were based on 20–45 glands from 4 HN, 6
DC, and 3 CF subjects. Error bars are SEM.
Evidence that SubP elevates [Ca2+]i. In a previous study, we presented evidence that CFTR-dependent synergy between VIP or
forskolin and low levels of carbachol resulted when activation of
Ca2+-activated K+ channels by carbachol increased the driving
force for anions through CFTR (19). In this
study, we used 3 methods to determine whether SubP might also elevate
[Ca2+]i and activate Ca2+-activated K+
channels: (a) We measured [Ca2+]i with Fura-2; (b) we compared
secretory responses to SubP in the presence and absence of the Ca2+
chelator BAPTA-AM; and (c) we also measured secretion in the presence or absence of
clotrimazole, an inhibitor of Ca2+-activated K+ channels
(Figure 7). In unstimulated cells,
[Ca2+]i was 70–120 nM. SubP increased
[Ca2+]i in 31 of 39 cells from 5 subjects by 139
± 33 nM (peak value). All 39 cells responded to carbachol with increases
in [Ca2+]i that were larger than those to SubP; the responses
to 1 and 10 μM carbachol were 187 ± 19 nM and 253
± 17 nM, respectively (Figure 7B).
We considered the possibility that gland cells that are unresponsive to SubP might be
a different cell type. To help differentiate serous and mucous cells in the dispersed
cell preparations, we used periodic acid–Schiff (PAS) staining and
observed a negative correlation between PAS reactivity and SubP responsiveness. For
SubP-responsive cells, 9 of 31 (29%) were PAS positive (contain mucus), while for
SubP-nonresponsive cells, 6 of 8 (75%) were PAS positive. This difference is
significant (P < 0.05, Fisher’s exact test).
Consistent with these measurements, in experiments with intact glands, BAPTA-AM (500
μM) reduced gland secretion stimulated by SubP by approximately 90%
(Figure 7C) and clotrimazole (25 μM)
reduced gland secretion stimulated by SubP by approximately 80% (Figure 7D). Although the number of experiments was small
for each of these 3 types of experiments, they mutually reinforced the interpretation
that SubP elevates [Ca2+]i and opens K+ channels.
Evidence that SubP stimulates gland secretion, in part, via elevating
[Ca2+]i. (A) Fluorescence changes in response to 10 μM SubP and 1
or 10 μM carbachol. Cell diameters in images are approximately 20
microns. (B) [Ca2+]i versus time for 6 cells
from images in A, measured in response to sequential pulses of 10
μM SubP and 1 and 10 μM carbachol. Fluorescence ratio, 340
nm/380 nm. (C) Mean response to SubP in presence or absence of
BAPTA-AM (500 μM); 4 experiments from 2 HN and 1 DC subjects
(16–20 glands). Error bars are SEM. (D) Mean response to
SubP in the absence and presence of clotrimazole (25 μM), which blocks
Ca2+-activated K+ channels (n = 4,
27–42 glands). *P < 0.05 versus SubP
responses. Error bars are SEM.
Pig glands responded more robustly than human glands to SubP. As previously reported (26, 30), porcine submucosal glands respond robustly
to SubP, with 1 μM SubP producing gland secretory rates equivalent to
those produced by 100 μM carbachol (26). However, in our experiments with humans, we observed much smaller
responses to SubP than to carbachol (e.g., Figure 2A). To determine whether there is a genuine species difference as opposed to
some difference between our procedures and those used previously (26), we directly compared secretory rates of
individual glands stimulated with carbachol or SubP in humans and pigs. The secretory
rates of single submucosal glands in pigs and humans exposed to 10 μM
carbachol were similar (data not shown), but secretory rates of glands exposed to 10
μM SubP were more than 10 times higher in pigs than in humans. In tissues
tested less than 7 hours from harvest, humans glands had a mean secretion rate of
0.29 ± 0.05 nl/min/gland (n = 8; 60 glands) compared
with a mean secretion rate in pigs of 4.02 ± 1.5 nl/min/gland
(n = 3; 28 glands; P < 0.001; Figure 8).
SubP stimulates airway gland secretion more strongly in pigs than in humans. A graph of (mean ± SEM) single gland secretion rates for 15 minutes
after 10 μM SubP application is shown (n = 3 pigs, 28
glands; n = 8 humans, 60 glands). Time from harvest to testing
was 4 ± 2 hours for pigs and 6 ± 2 hours for humans.
***P < 0.001.
Features of gland responses to SubP studied with DIC microscopy. To determine the effects of SubP on cells within the glands, single submucosal glands
were microdissected from airway mucosa and mounted as described in the methods
section for observation with DIC optics, using time-lapse digital imaging. Under our
typical conditions (×40 water immersion lens), the field usually
contained only 1 or 2 serous acini or mucous tubules out of the more than 100 that
make up a gland (ref. 36 and see below).
Responses of tubules or acini were monitored quantitatively by measuring changes in
the outer diameters and lumens of the tubules and acini, which reflect cell volume
changes, as well as qualitatively by observing movement of particles in the mucus
when visible. Responses of HN and DC subjects were similar and were combined for
comparison with CF glands. When SubP (1–10 μM) was superfused
over the glands, we observed rapid shrinkage of acinar serous cells by
15%–50% of their height and 6% of their calculated volume, with an
associated increase in lumen volume. Mucous cells in tubules from control subjects
did not change volume significantly (increase of 2% ± 5%;
n = 4), although the response was quite variable (range, increase of
17% to decrease of 7% calculated cell volume change).
The mean peak change in normalized cell height was 26% ± 14% for 7
control subjects (3 DC and 4 HN subjects) where clear images were obtained following
exposure to SubP. (The mean cell heights in control cells before application of SubP
were serous, 21.2 ± 2.0 μm and mucous, 20.6 ± 2.7
μm.) We also observed the following features: (a) increased flow of mucus
from the serous acini and in tubules and ducts, as evident by the movement of
particles within the mucus; (b) small and transient myoepithelial cell contractions;
(c) persistence of these effects in the presence of 1 μM atropine (which
abolished the response to 1 μM carbachol; data not shown). Examples of
responses in 1 HN and 1 DC gland are shown in Figures 9 and 10, in which the increase in
lumen volumes is plotted. Twelve glands from eleven control subjects were studied,
and all responded to 1 or 10 μM SubP except for 1 gland from a
56-year-old patient with chronic obstructive pulmonary disease (COPD).
Isolated HN gland exposed to SubP shows cell shrinkage, lumen volume
expansion, and fluid secretion when monitored with DIC. (A) One lobe of an isolated human gland. Scale bar: 250
μm. (B) Serous acini and mucous tubules just prior to
stimulation. (C) Same gland structures 100 seconds after addition of
10 μM SubP plus 1 μM phosphoramidon. Scale bar: 25
μm (B and C). (D) Plots of
volumes of serous and mucous lumens as a function of time and stimulation
(indicated with gray bar; see Methods).
Isolated DC gland exposed to SubP shows cell shrinkage, lumen volume
expansion, and fluid secretion when monitored with DIC. The gland is from a subject with COPD. (A) Serous acini and mucus
tubules just prior to stimulation. (B) The same structures at peak of
response to 10 μM SubP plus 1 μM phosphoramidon
(phosphor). Scale bar: 20 μm (A and B).
(C) Plots of volumes of serous and mucous lumens as a function of
time and stimulation (indicated with gray bar; see Methods).
In contrast with the reliable and stable acinar cell volume loss and lumen expansion
observed in airway glands from control subjects in response to SubP, the gland cells
in 3 of 4 CF subjects showed what appeared to be transient or sustained
expansion of cell height by 5%–15% and a concomitant
decrease of lumen volume (Figure 10, A–C). However, as described in Methods, when linear
changes in the 2-dimensional optical slices were used to model volume changes in the
3-dimensional tubules of the glands, the apparent “reversed”
responses of the CF glands to SubP were seen to arise from the combination of a
smaller, slower, and transient loss of cell volume and a superimposed transient
contraction of the myoepithelial cells, resulting in lumen closure (Figure 11). The gland from 1 CF subject (CF48, who also
responded when tested with the oil layer method; Supplemental Figure 1) responded to
SubP with cells shrinkage, lumen expansion, and mucus flow from the acinus, but
unlike control glands, this response was transient in the continued presence of SubP
(Figure 11, D–F).
Aberrant responses in CF gland acini to SubP. (A) Acinus and tubule just prior to stimulation. (B)
Same structures at peak response to 10 μM SubP plus 1 μM
phosphoramidon. The acinus was outlined and an arrow was drawn spanning cell
height in the control state, and these were then copied onto the image of the
acinus at peak stimulation to show changes. (C) Plots of lumen
volumes assuming them to be cylinders (see Methods) for 4 CF subjects stimulated
with SubP. A decrease in lumen volume was observed in 3 of 4 CF
subjects stimulated with SubP; these subjects did not secrete mucus when assayed
with oil layer method. CF48 responded with a normal increase in lumen volume but
with a much shorter time course than controls. The gray bar indicates the period
of SubP stimulation. (D–F) Lumen expansion
of CF48. Images are just before (D), at peak (E), and at
minimal lumen volume (F) (which was maintained in this gland) in
response to 10 μM SubP plus 1 μM phosphoramidon and 10
μM atropine. Scale bar: 20 μm.
The DIC results help clarify our finding that many glands were
“nonresponsive” to SubP when we measured mucus secretion with
the bubble method. All human airway glands that we tested responded to SubP, but in
some glands, the stimulated secretion rates are not sufficient to result in
measurable flow of mucus from the gland duct opening. This may occur because of
physical capacitance within the glands (i.e., lumen dilation without mucus exit),
which could be exacerbated if there is a mismatch between fluid and macromolecular
secretion, leading to higher viscosity.
For time-lapse sequences for selected control and CF gland responses to SubP, see
Supplemental Videos 1–3.
The present results in humans and prior results in mice (18, 31) establish that mucosal chili
oil activates airway submucosal glands, at least in part via a pathway that releases
SubP, and that in these 2 species, SubP stimulates the glands by a CFTR-dependent
mechanism. In humans, SubP must work by at least one pathway that is not involved with
the elevation of cAMP, because when SubP is added to saturating concentrations of
forskolin, it produces a synergistic increase in secretion. One possible mechanism for
such synergy would be elevation of [Ca2+]i by SubP and subsequent
activation of basolateral Ca2+-activated K+ channels, as suggested
by the data in Figure 7. The small secretory
response to SubP, its near-total dependence upon CFTR, and its synergy with forskolin
suggest that, unlike carbachol, it does not activate apical Ca2+-activated
Cl– channels. We do not know how this occurs, but it also
occurred in previous synergy experiments with low-dose carbachol (19). Possible reasons for this are that the elevation of
[Ca2+]i is subthreshold for Ca2+-activated
Cl– channels and that the [Ca2+]i
elevation is spatially restricted in the cells. The experiments measuring
[Ca2+]i with Fura-2 support the first interpretation.
Activation of CFTR-dependent secretion by classical [Ca2+]i
elevating agents like ACh also occurs in the intestine, in which
Cl– secretion stimulated by ACh is completely lost in CF (37–39). In the intestine, Cl– secretion stimulated by
Ca2+-elevating agents depends upon the elevation of [cAMP]i,
secondary to prostaglandin release, because it can be blocked with indomethacin in human
(40) or mouse colon (41) and it is strongly synergistic with PGE2 in the
guinea pig colon (42). In airway submucosal
glands, it is also possible that SubP induces an elevation of [cAMP]i in
gland cells, but we have no direct evidence of this.
In the intestine, the CFTR dependence of Ca2+-mediated secretion can be
observed clearly, because the intestinal cells typically lack Ca2+-activated
Cl– channels (43).
However, human and mouse airway submucosal glands certainly contain
Ca2+-activated Cl– channels. Carbachol has been shown
to elevate [Ca2+]i in mouse nasal gland acinar cells (28), and bumetanide-sensitive and niflumic
acid–sensitive fluid secretion in response to carbachol is robust in CF
human glands (present results and ref. 7) and in
CF mouse glands (18), yet SubP, unlike carbachol,
failed to activate these channels (or activated them only weakly; see below).
The hypothesis that SubP stimulates serous cell fluid secretion by activating
basolateral Ca2+-activated K+ channels but not apical
Ca2+-activated anion channels is the same hypothesis put forth to explain
CFTR-dependent synergy between low levels of cholinergic and VIP stimulation (19). Thus, SubP, which by itself is a much weaker
secretagogue in human and mouse glands than carbachol, appears to produce effects
similar to those of low concentrations of carbachol. This hypothesis might account for
the rare secretory responses of human CF glands to SubP: in some glands the
Ca2+ signal might be strong enough to activate some apical
Ca2+-activated anion channels. Regardless of the cellular mechanism, the main
point of our experiments is that SubP-mediated fluid secretion from human airway glands
is CFTR dependent and is synergistic with VIP or forskolin.
Distinguishing types of gland secretion. When considering mucus secretion by submucosal glands, it is important to distinguish
at least the following 3 components that make up bulk mucus: mucin secretion,
non-mucin protein secretion, and electrolyte-driven fluid secretion. The bubble
method used here measures total mucus volume, which is primarily water. This
contrasts with most early studies of airway gland secretion that relied solely on
markers of macromolecular secretion. Thus, Rogers et al. measured fucose, hexose, and
proteins in the apical fluid of Ussing chambers as biochemical markers of mucus
secretion from bronchial explants from CF and non-CF tissues (44). They found similar amounts of markers in the basal state.
However, after stimulation with methacholine, terbutaline, or SubP, they observed
large increases for each marker in control tissues, but in CF tissues, the increases
were significantly smaller for all 3 agonists. As they recognized, the reduced levels
of macromolecular components in the apical fluid is consistent either with fewer
macromolecules being secreted or with CF mucus being “more viscid than
normal and . . . less easily collected” (44). Much evidence now favors the latter interpretation. The loss of CFTR
inhibits anion-mediated fluid secretion, so that the normally rapid and extensive
expansion of mucin granules is slowed and constrained, leading to a more densely
entangled and hence more viscous network (5).
Impacted secretions with high solids and viscosity were produced in pig airway glands
by inhibiting anion transporters (14–16), and undispersed
Paneth cell granules accumulate in the intestinal crypts of CFTR-knockout mice (45). In this regard, it was recently proposed
that HCO3– plays a crucial role in enabling mucin
granule expansion by competing away the charge shielding Ca2+ and
H+ ions that are required to compact mucins in granules. The known
deficiency of HCO3– secretion in many CF organs
could thus contribute to altered mucus properties (46). Thus, although our measurements indicate that the volume of
carbachol-stimulated mucus is normal in CF, it arises from hypertrophied glands, and
its properties may well be changed so that it is less easily cleared or collected
The present results illustrate the utility of combining DIC observations of cell
responses in intact, isolated glands with measurements of volume secretion by
individual glands. In DIC experiments, SubP caused HN or DC acinar serous cells to
lose volume for as long as SubP was present, while, concomitantly, the gland lumens
expanded and mucus flowed from the acini and along the tubules (see also ref. 4). The loss of cell volume coupled with lumen
expansion and mucus flow is a marker of electrolyte-driven fluid secretion (e.g.,
ref. 48). Myoepithelial cell contractions were
small and transient, indicating that sustained or rhythmic contractions are not
required for mucus secretion.
In CF glands, DIC markers of fluid secretion stimulated by SubP were absent in 3 of 4
subjects. Importantly, all CF glands responded transiently to SubP, proving that
receptors are still present, but the transient cell volume loss was not accompanied
by other markers of fluid secretion, and the brief myoepithelial contractions
actually reduced the lumen volumes in the CF cells.
Species differences. We found that SubP was many fold more efficacious in pig versus human glands. In this
regard, human glands were more like mouse glands than pig glands, because mouse
glands also show a much smaller response to SubP than they do to carbachol (18, 31).
The approximately 10-fold increased efficacy of SubP for stimulating mucus secretion
from pig versus human glands is consistent with a much greater density of mucosal
SubP innervation in the pig bronchial mucosa, which has a profuse apical plexus of 1
μm diameter axons (~2,000 terminations per mm2, 94% of which
are immunoreactive for SubP) (49). Although a
similarly dense neural plexus was observed in human bronchial tissues, SubP
immunoreactivity was barely detectable in it (49).
We hypothesize that the density of SubP receptors is higher in pig glands than it is
in human glands. If true, this could result in a larger increase in
[Ca2+]i within the pig gland cells. This would be expected
to increase secretion through active CFTR and also might be sufficient to recruit
apical Ca2+-activated Cl– channels. If the latter
is true, a larger component of SubP-mediated secretion may be spared in CFTR-knockout
pigs than in CF humans. Whether this will have relevance for the development of
airway disease in the CFTR-knockout pig can now be tested directly (50, 51).
This species difference is the largest we have yet observed between humans and pig
airway glands and is under further investigation (M. Khansaheb, unpublished
observations). However, even more dramatic differences have been described for SubP
effects in pigs and humans: application of SubP to the iris sphincter muscle of pigs
increased inositol triphosphate (IP3) accumulation and contracted the
muscle, while in humans, it did not increase IP3 or cause contractions but
instead increased cAMP production (52).
Relevance to CF airway disease. The ability of mucosal irritants to evoke gland secretion in control but not CF
tissues may be one reason that low levels of local pathogens are not quickly cleared
from CF airways. The hypothesis that CF airway infections are related to abnormal
mucus properties and clearance is increasingly attractive (reviewed in refs. 5, 9, 53). Because most upper airway mucus arises from
glands, it is critical to define the role of CFTR in mucus secretion by airway
glands. The present study shows that SubP, like VIP, activates secretion via a
process that is mainly CFTR dependent, and yet, the 2 agonists are synergistic and
their combined effects are also CFTR dependent. We remain ignorant of how these
processes are employed in normal airways. Normal breathing in quiescent animals
produces a level of neural tone in airway parasympathetic nerves, (54) and we speculate that glands are stimulated
at low levels by such input. Our prior results show that CFTR plays a proportionately
larger role during low levels of gland fluid secretion (19). The present results reinforce that conclusion, and may also
be interpreted to suggest that local irritants, acting through axonal reflex rather
than central reflex pathways, also stimulate CFTR-dependent gland secretion. If this
reasoning is correct, then hyposecretion of anion-mediated fluid by CF airway glands,
both in the “basal” state and in response to minor local
irritants, would be expected to hamper the initial formation of the mucus gel (5, 7, 10–24, 46), thus limiting the normal
dispersion of antimicrobial-rich mucus over the airway surface and leading instead to
accumulation within glands (14–16) and, via
tethering to gland orifices, to mucus plugging of smaller airways. These consequences
would be expected to diminish the innate mucosal defenses of CF airways and set the
stage for chronic infections.
Human airway preparations. These studies were approved by the Institutional Review Boards of Stanford University
and Yonsei University. At Stanford University, human bronchial tissues were obtained,
after prior written consent, immediately following lung transplants. Surgical scrap
tissues were obtained from donor tracheae at Stanford University. At Yonsei
University, tracheotomy flaps from subjects without lung disease were obtained after
prior written consent.
Data were obtained from 32 HN donor tracheae, 5 HN tracheotomy flaps, 12 CF patients,
and 21 DC patients, who had transplants for diseases other than CF. The HN donor
tracheae were obtained at each transplant as surgical scrap (donor lungs are
transplanted one at a time with anastomosis at the level of the mainstem bronchi).
The 21 DC lungs were from 10 patients with COPD, 4 patients with pulmonary fibrosis,
3 patients with primary pulmonary hypertension, 3 patients with interstitial lung
disease (ILD) (1 patient with ILD/sarcoidosis, 1 patient with ILD/neurofibromatosis,
and 1 patient with ILD/unknown), and 1 patient with idiopathic pulmonary
hemosiderosis. Genotypes were available for 8 of the CF subjects; 5 were
ΔF508 homozygous, with 1 of each of the following:
ΔF508/N1303K, G542X/W1282X, and 406-1 G->A/H119Y. Subject
characteristics for gland secretion experiments are given in Table 1 and for DIC experiments are given in Table
2. One CF and two HN patients were studied
only with DIC, all others were also used in gland secretion experiments. The DC
subjects used for DIC comprised 3 COPD patients and 1 familial pulmonary fibrosis
subject. Tissues used for measurements of [Ca2+]i in acinar
cells were tracheotomy flaps obtained from patients without pulmonary disease
(n = 5, 53 ± 18 years).
Subject characteristics for isolated gland DIC measurements
All tissues were transported to the laboratory in cold Physiosol solution (Abbott
Laboratories) and were then transferred to ice-cold Krebs-Ringer bicarbonate buffer
(KRB), bubbled with 95% O2–5% CO2, in which they
were maintained until use, usually within 24 hours. The KRB composition was 115.0 mM
NaCl, 2.4 mM K2HPO4, 0.4 mM KH2PO4, 25.0
mM NaHCO3, 1.2 mM MgCl2, 1.2 mM CaCl2, 10.0 mM
glucose, and 1.0 μM indomethacin. KRB was made to 90% volume, and the
osmolarity was measured with a Wescor 5500 Vapor Pressure Osmometer. Distilled water
was added to adjust the osmolarity to 290 ± 5 mOsm. The pH was verified
to be 7.4 (Orion 420A pH meter) after bubbling with 95% O2–5%
Pig tracheae. Pig tracheae were obtained from fresh carcasses of 3 Juvenile Yorkshire pigs of
either sex, weighing 35~50 kg, following acute experiments carried out for other
purposes. Procedures for care and euthanization of pigs were approved by the
Administrative Panel on Laboratory Animal Care (Stanford University’s
Institutional Animal Care and Use Committee); no pigs were sacrificed specifically
for the present experiments.
Optical measurement of mucus secretion rates (mucus bubble method). To prepare tissues for optical recording of mucus secretion rates by individual
glands, a piece of ventral trachea or bronchus of approximately 0.5 cm2
was pinned mucosal side up and the mucosa with underlying glands was dissected from
the cartilage and mounted in a 35-mm diameter, Sylgard, lined plastic Petri dish (Dow
Corning Corporation), with the serosa in the bath (~1 ml volume) and the mucosa in
air. The tissue chamber was maintained at
35°C–37°C and high humidity, using either a
Sensortek S-4 Peltier effect TC-102 Temperature Controller and Fisher-Milligan Gas
Washer or a thermistor-controlled LU-CB1 warming chamber and humidifier (Medical
Systems Corp.). The tissue surface was cleaned and blotted dry with cotton swabs and
further dried with a stream of gas, after which 20–30 μl of
water-saturated mineral oil was placed on the surface. The tissue was warmed to
37°C at a rate of approximately 1.5°C
min–1 and continuously superfused with warmed, humidified 95%
O2–5% CO2. Pharmacological agents were diluted
to final concentration with warmed, gassed bath solution and were added to the
serosal side by complete bath replacement. Bubbles of mucus within the oil layer were
visualized by oblique illumination and digital images were captured either with the
macro mode of a Nikon digital camera or by mating a digital camera to 1 ocular of a
Wild stereomicroscope. Each image contained an internal reference grid to compensate
for any minor adjustments in magnification made during the experiment. Stored images
were analyzed either by direct measurement or with ImageJ software
(http://rsb.info.nih.gov/ij/). Mucous volumes were determined from the size of the
spherical bubbles; bubbles that were not approximately spherical were omitted from
secretion rate analyses and are given as nl/min/gland. Details of these methods are
given in ref. 33.
Secretion rates for individual glands vary over a many fold range. For synergy
experiments, variations in gland secretion rates to forskolin, SubP, and the
combination were minimized by normalizing the rates to the peak secretion rates for
carbachol in those same glands tested at the end of the experiment.
Estimating the proportion of responding glands. In past experiments, we found carbachol to be the most efficacious agonist for
stimulating mucus secretion of those we tried with human glands, and at high (10
μM) concentrations, carbachol continued to stimulate secretion in CF
glands. Therefore, we used responsiveness to a 5-minute exposure to 10 μM
carbachol at the end of experiments to estimate the number of viable glands in each
preparation; the number of glands responding to SubP was then divided by the number
responding to carbachol to estimate the proportion of SubP-responsive glands.
Fura-2 measures of [Ca2+]i. Tracheal submucosal glands were isolated using a Wild zoom binocular dissecting
microscope and transferred to 2 ml of bicarbonate-buffered Ringer’s
solution (125.0 mM NaCl, 0.4 mM KH2PO4, 1.6 mM
K2HPO4•3H2O, 1.0 mM MgCl2,
5.0 mM glucose, 10.0 mM Na-acetate, 2.0 mM glycine, 1.0 mM
α-ketoglutarate, 2.0 mM CaCl2, 25.0 mM NaHCO3)
gassed with 95% O2/5% CO2. After 7 minutes of incubation in
this buffer solution, containing 2 mg/ml of Collagenase NB 4 Standard Grade (SERVA)
and 2 mg/ml of Trypsin Inhibitor (Sigma-Aldrich) at 37°C, submucosal
glands were agitated gently using a fire-polished, wide-bore (1–2 mm)
Pasteur pipette. After sedimentation the gland cells were washed twice with
enzyme-free bicarbonate-buffered Ringer’s solution. The isolated acinar
fragments were seeded onto glass cover slips (22 × 22 mm) coated with
poly-l-lysine and were incubated for 30 minutes in physiologic salt
solution (140 mM NaCl, 5 mM KCl, 1 mM MgCl2, 1 mM CaCl2, 10 mM
HEPES, and 10 mM glucose), containing 5 μM fura-2, AM (Teflabs) in the
presence of Pluronic F-127 (Invitrogen) to enhance dye loading.
Fura-2–loaded cells were mounted in a flow chamber on the stage of an
inverted microscope (Nikon) for imaging; temperature was maintained at
37°C. The cells were illuminated at 340 nm and 380 nm, and the emitted
fluorescence at 510 nm was collected with a CCD camera and analyzed using the
MetaFluor system (Universal Imaging). The fluorescence ratio (340 nm/380 nm) was
taken as the measure of [Ca2+]i and fluorescence images were
obtained at 2 second intervals.
[Ca2+]i was calculated by using following equation (55): [Ca2+]i =
– R)]β, where Kd
is the dissociation constant of fura-2 for Ca2+ (224 nM at
37°C), r is the ratio of fluorescence of the sample at
340 and 380 nm, Rmax represents the saturated
Ca2+ level determined by addition of 10–5 M
ionomycin, Rmin represents the free Ca2+ level
controlled by 5 mM EGTA, and β represents the ratio of fluorescence at
380 nM in no Ca2+ and saturating Ca2+.
DIC imaging of isolated airway submucosal glands. Tissues containing glands were isolated as described above and then were further
microdissected to prepare relatively clean but intact glands for optimal imaging.
When judged suitable for imaging, the glands were transferred to microperfusion
chambers on the stage of an upright Nikon Eclipse E600FN Series Microscope equipped
with DIC and epifluorescence. They were continuously perfused with Krebs, gassed with
95% O2/5% CO2 via a pressurized, 8-chamber, solenoid actuated
perfusion system (Automate). Temperature was maintained at
35°C–37°C using a TS-4 Peltier effect temperature
controller that warmed the inflow tubing and chamber. For digital imaging, the
microscope was fitted with a Retiga-1300, cooled, 12-bit, color Bayer mosaic CCD
camera (QImaging) with RGB Liquid Crystal Color Filter Module (Spectra). The camera
is interfaced with a computer running Compix Image capture and analysis software. All
human glands were tested within 15 hours of harvest (12 ± 3 hours for DC
subjects; 10 ± 5 hours for HN subjects; 12 ± 2 hours for CF
For overview images of the gland, we used an ×4 objective. For detailed
studies of cellular changes in the acini, tubules, and ducts of glands, glands were
imaged with an ×40 water immersion lens (numerical aperture, 0.8; 2-mm
working distance) and time lapse digital imaging was used to monitor changes with
frame rates of 2–30 s–1. At this magnification,
the field and focal plane are sufficiently restricted, so that it is usually
necessary to pick a single duct, tubule, or acinus for optical imaging.
Serous and mucous cells were distinguished by 2 criteria: location and physical
appearance. Cells that were granular when viewed with DIC and that were only seen in
the acini of the gland were scored as serous cells; these cells usually appeared to
have a smaller apical surface than basolateral surface. Cells in the tubules
downstream from the acini appeared smooth when viewed with DIC, and unlike the
tapering serous cells, they were typically rectangular when viewed from the side;
these were scored as mucous cells. Usually an abrupt transition occurred between the
2 types of cells, sometimes indicated by a constriction. On occasion, we saw acini
that appeared to be a continuation of the mucous tubule, with no serous cells that we
could discern; these mucous acini have been described previously (36).
The DIC method produces a thin optical slice through an acinus and/or tubule. We
obtained longitudinal optical sections whenever possible to allow observation of
linear flow of mucus particles as an independent, qualitative measure of secretion.
We expected, on the basis of much other work (e.g., 28, 29), that secreting cells might
show volume changes. These were readily observed and could be quantified by making
the simple assumptions (based on numerous observations) that tubules are essentially
cylinders and acini are their spherical ends (perhaps slightly enlarged); the open
portion of the cylinder is the lumen. The longitudinal optical section of the tubules
and acini allowed us to measure their outer diameter, lumen diameter, and length
(from specified landmarks) before, during, and after stimulation at a number of
points along the tubule/acinus. Measurements were made on high-resolution,
precalibrated digital images, captured every 2–30 seconds using Compix
analysis software. Measurements were only made when 2 criteria were satisfied: both
the outer edges of the tubule/acinus and the corresponding edges of the lumen had to
be clearly visible, and (assuming they are cylinders), their initial diameters were
maximized by through-focusing, so that we knew we were measuring the center of the
acinus/tubule and the lumen.
We used the following rationale for interpreting our data. A transection through a
duct, tubule, or acinus would appear as an annulus with an area occupied by cells =
π × (outer radius2 – lumen
radius2). If our assumption of cylindrical shape was valid, then we
measured these radii and so could compute the volume of cells in the annulus for an
arbitrary length of tubule (i.e., the thickness of the annulus), which we arbitrarily
set to 20 μm. It was a consistent finding that this volume decreased
during stimulation, because the outer radius did not change (or became smaller),
while the inner radius (lumen) increased. We refer to this as a net cell
volume loss, because we can’t know if all of the cells in the
annulus respond identically; but if some don’t change or increase their
volume, then correspondingly larger decreases must occur in other cells to give the
aggregate decrease in volume we observed.
These measurements capture only a fraction of the dynamics of gland behavior during
stimulation; therefore, video files have been provided as supplemental online
material (Supplemental Videos 1–3).
Reagents. Compounds (from Sigma-Aldrich, Teflab, Invitrogen, and SERVA) were made fresh or
maintained at –20°C. Stock solutions of SubP, phosphoramidon
(a metalloproteinase inhibitor that helps prevent breakdown of SubP by
endopeptidase), carbachol, VIP, atropine (inhibitor of muscarinic receptors), and
clotrimazole were dissolved in sterile distilled water; indomethacin (inhibitor of
prostaglandin release) was dissolved in ethanol; and forskolin and BAPTA-AM were
dissolved in DMSO. All were diluted 1:1,000 with bath solution (except indomethacin,
1:10,000) immediately before use at the concentrations indicated. The highest DMSO
concentration in our experiments was 0.20%, and the highest ethanol concentration was
0.01%. We tested 0.5% DMSO as vehicle alone with no effect. To stimulate mucosal C
fibers, we used a commercially available hot chili pepper oil that contained
capsaicin and other irritants (Melina’s; Pepper Mill Imports).
Statistics. Data are mean ± SEM. Unpaired, 2-tailed Student’s
t test was used to compare the means of different treatment groups
unless otherwise indicated. Fisher’s exact test was used for 2
× 2 contingency tables. A 1-way ANOVA with Tukey-Kramer post-hoc test was
used for comparisons of 3 or more data sets. Any difference was considered to be
significant when P was less than 0.05. Curves were fit with Origin
software (OriginLab Corporation) using a sigmoid function.
View Supplemental data
View Supplemental video 1
View Supplemental video 2
View Supplemental video 3
Research was supported by NIH (DK-51817) (to J.J. Wine), the Cystic Fibrosis Foundation,
and Cystic Fibrosis Research Inc. J.Y. Choi and M. Khansaheb were recipients of
Elizabeth Nash Fellowships. We are very grateful to John Hanrahan and Juan Ianowski,
with whom we initiated the study of SubP on gland secretion in mice and whose insights
have greatly helped the present work. We thank Tony Nguyen and Kim Tran for valuable
technical assistance and for tissue harvesting, Marlene Wine and Kim Tran for help
obtaining consents, Lisa Levin and Jackie Zirbes for help with patient information, and
Jennifer Lyons for helpful discussions. We especially thank the patients and their
families, whose donations of tissues were essential for this research.
Conflict of interest: The authors have declared that no conflict of
Nonstandard abbreviations used: ACh, acetylcholine;
[Ca2+]i, intercellular Ca2+ concentration;
[cAMP]i, intercellular cAMP concentration; CF, cystic fibrosis; COPD,
chronic obstructive pulmonary disease; DC, disease control; DIC, differential
interference contrast; HN, human normal; SubP, substance P; VIP, vasoactive
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