Developmental regulation of intestinal angiogenesis
by indigenous microbes via Paneth cells
Thaddeus S. Stappenbeck, Lora V. Hooper, and Jeffrey I. Gordon*
Department of Molecular Biology and Pharmacology, Washington University School of Medicine, St. Louis, MO 63110
Contributed by Jeffrey I. Gordon, October 7, 2002
The adult mouse intestine contains an intricate vascular network.
The factors that control development of this network are poorly
understood. Quantitative three-dimensional imaging studies re-
vealed that a plexus of branched interconnected vessels developed
in small intestinal villi during the period of postnatal development
that coincides with assembly of a complex society of indigenous
gut microorganisms (microbiota). To investigate the impact of this
environmental transition on vascular development, we compared
the capillary networks of germ-free mice with those of ex-germ-
free animals colonized during or after completion of postnatal gut
formation. The developmental program can be restarted and
completed within 10 days after colonization with a complete
microbiota harvested from conventionally raised mice, or with
Bacteroides thetaiotaomicron, a prominent inhabitant of the nor-
mal mouse/human gut. Paneth cells in the intestinal epithelium
secrete antibacterial peptides that affect luminal microbial ecol-
ogy. Comparisons of germ-free and B. thetaiotaomicron-colonized
transgenic mice lacking Paneth cells established that microbial
regulation of angiogenesis depends on this lineage. These findings
reveal a previously unappreciated mechanism of postnatal animal
development, where microbes colonizing a mucosal surface are
assigned responsibility for regulating elaboration of the under-
lying microvasculature by signaling through a bacteria-sensing
small intestine ? gnotobiotics ? ecology ? symbiosis
finger-shaped projections (villi). The gut epithelium undergoes
constant renewal throughout postnatal life. This process is
fueled by one or more long-lived active multipotent stem cells
rise to four lineages, three of which (enterocytic, goblet, and
enteroendocrine) complete their differentiation as they migrate
out of crypts up onto adjacent villi. In contrast, members of the
fourth lineage (Paneth cell) complete their differentiation at
the crypt base (3, 4). Paneth cells are critical components of the
intestine’s innate immune system; they secrete a wide variety of
antimicrobial peptides and proteins into the gut lumen (5, 6).
The intestinal epithelium is uniquely positioned to receive and
transduce signals from two vastly different cellular populations.
It sits at the interface between a subepithelial ensemble of
mesenchymal fibroblasts, immune and endothelial cells, and a
vast community of indigenous microorganisms (the microbiota)
that reside in the gut lumen (7). Endothelial cells in the villus
distribution of absorbed nutrients.
Collectively, the microbiota can be viewed as a metabolically
active ‘‘organ’’ that provides functions beneficial for both mi-
crobes and host. For example, by supporting assembly of a
society of resident microbes with the capacity to break down
carbohydrate polymers in the distal intestine (e.g., refs. 9–16),
mice (and humans) did not have to evolve their own repertoire
of glycosylhydrolases to cleave the wide variety of linkages that
exist in dietary polysaccharides. In this arrangement, the host is
he mucosal surface of the adult mouse and human small
able to extract nutrient value from an otherwise poorly used
dietary substrate, while microbes are provided with a niche
where they can gain access to abundant, readily fermentable
The idea that our resident microbial communities are impor-
tant contributors to normal mammalian physiology and health is
now developing a broader appreciation of the wide range of host
functions that are regulated by symbiotic microorganisms
(reviewed in refs. 18–21). In the present study, we examine the
interactions between the gut microbiota, the small intestinal
epithelium, and the villus’ mesenchymal microvascular network.
We show that the microbiota plays a key role in constructing this
microvascular network, and that this regulation depends on a
central component of the gut’s innate immune system: the
Paneth cell. These findings reveal a previously unappreciated
pathway for control of angiogenesis, and emphasize the impor-
tance of considering features of postnatal mammalian develop-
ment as manifestations of mutually beneficial partnerships with
Materials and Methods
Mice. All experiments with mice were performed using protocols
approved by our institutional animal studies committee. Germ-
free mice belonging to the NMRI inbred strain were maintained
in plastic gnotobiotic isolators (22) under a 12-h light cycle, and
given free access to autoclaved water and chow (B & K Uni-
versal, East Yorkshire, U.K.). We re-derived FVB/N CR2-tox176
transgenic mice (23) as germ-free using protocols described in
ref. 22, and maintained the pedigree by crosses to germ-free
nontransgenic littermates. Animals were genotyped by PCR
assays of tail DNA (23). Genotypes were confirmed at the time
of sacrifice by documenting Paneth cell loss through staining
small intestinal sections with indocarbocyanine (Cy3)-labeled
Ulex europeaus agglutinin-1 (ref. 24; Sigma).
Colonization of Germ-Free Animals. Mice were inoculated with
either mid-log phase cultures of Bacteroides thetaiotaomicron
(type strain VPI 5482), or an intestinal microbiota harvested
from the distal third of the small intestines and cecums of
age-matched, conventionally raised, specified pathogen-free an-
imals (22). Mice were killed 7, 10, or 30 days after inoculation.
Animals with ?107colony forming units per ml luminal contents
in their distal small intestines were used to evaluate the villus
Quantitative Analysis of the Villus Microvasculature. All mice (con-
ventionally raised, germ-free, and ex-germ-free) were killed at
the same time of day (12:00 p.m.). Animals were anesthetized by
metofane inhalation for 5 min, and then injected with 100 ?l of
a 2 mg/ml solution of high-molecular-weight (2,000 kDa) fluo-
rescein isothiocyanate (FITC)-labeled dextran (Sigma). A 30-
Abbreviation: Pn, postnatal day n.
*To whom correspondence should be addressed at: Department of Molecular Biology and
Pharmacology, Washington University School of Medicine, Box 8103, 660 South Euclid
Avenue, St. Louis, MO 63110. E-mail: email@example.com.
November 26, 2002 ?
vol. 99 ?
no. 24 ?
gauge needle, attached to a 1-ml syringe, was used to infuse this
material over a 10-s period into the retro-orbital plexus. Animals
were killed 3 min later. The small intestine was removed and
measured without stretching. An 8-cm segment, spanning the
junction of the middle and distal thirds of the small bowel, was
excised, perfused with fixation solution containing 0.5% para-
formaldehyde, 15% picric acid, and 0.1 M sodium phosphate
buffer (pH 7.0), opened with an incision along its cephalocaudal
axis, pinned on black wax, and shaken gently at 4°C for 12 h in
fixation solution. Specimens were rinsed in ice-cold PBS (three
washes, 5 min each), followed by a 3-h incubation in 10%
sucrose/PBS (4°C) and an overnight incubation in 20% sucrose/
10% glycerol/PBS (4°C).
Villi were examined using two different methods. In the first
method, 2.5-cm-long portions of the intestinal segment were
frozen in OCT compound, and 120-?m-thick sections were cut
at room temperature in the dark, followed by rehydration in
ice-cold PBS (1 min) and an overnight incubation at 4°C in 3%
deoxycholic acid (Sigma). Sections were rinsed two times in
water (5 min per cycle, room temperature), and one time in PBS
(5 min, room temperature) to remove residual deoxycholic acid.
Sections were stained with Syto61 (Molecular Probes; 1:1,000
dilution in PBS) for 1 h at room temperature, followed by three
PBS washes (5 min per cycle, room temperature). Sections were
mounted in 50% glycerol/PBS and stored at 4°C before viewing.
In the second method, whole-mount preparations are used
rather than cryosections. A 1-cm portion of the fixed intestinal
segment was washed in PBS and stained with Syto61 exactly as
in the first method. The stained whole-mount preparation was
placed on a glass slide (villus side up), and 100 ?l of a solution
of 50% glycerol/PBS was added, followed by a coverslip.
Cryosections and whole mounts were viewed with an LSM 510
confocal microscope (Zeiss) and scanned at 5-?m-thick inter-
vals. Scans were projected in three dimensions by taking 12–16
serial images, aligning them at 7–10° intervals, and compiling/
rotating them about the y axis by using LSM 510 software (Zeiss).
The capillary network in the upper third of villi was quantified
by counting the number of Syto61-positive nuclei per window as
visualized from a fixed perspective of the three-dimensional
reconstruction (a window was defined as an open area bounded
by capillaries; n ? 20–25 villi per mouse; n ? 3 animals per
group). Mean values ? SD of nuclei per window were calculated
for each mouse and mean values ? SEM computed for each
group. The statistical significance of differences observed be-
tween groups was assessed using Student’s t test.
PHOTOSHOP (Version 5.0, Adobe Systems, Mountain View,
CA) was used to create two-dimensional images from rotating
three-dimensional images. These two-dimensional images are
shown in Figs. 1–3.
A Quantitative Assay for Visualizing Development of the Microvascu-
lature in Three Dimensions Within the Mouse Small Intestine. Two-
dimensional views are not adequate for characterizing the
organization of the villus microvasculature (Fig. 1 A and B).
Therefore, we developed a quantitative method for visualizing
blood vessels in three dimensions. This method employs retro-
orbital injection of a solution containing high-molecular-weight
FITC-conjugated dextran, followed by confocal microscopy of
villi present in either whole-mount preparations of small intes-
tine or 120-?m-thick cryosections. Epithelial nuclei were labeled
with Syto61, and the density of the microvascular network was
quantified by counting the number of stained nuclei overlying
the open spaces or ‘‘windows’’ framed by the underlying inter-
connected capillaries. The higher the density of capillary
branches and connections, the smaller the average number of
nuclei contained per window. By assembling serial confocal
prepared from the junction between the middle and distal thirds of the small
from the same animal as in A, but stained with H&E and antibodies to the
microscopic scan of a 120-?m-thick cryosection. The capillary network is stained
with FITC-tagged high-molecular-weight dextran (green), and epithelial nuclei
with Syto61 (red). To view a three-dimensional rotating image of a compiled set
information on the PNAS web site, www.pnas.org, or go to http://gordonlab.
wustl.edu/vasculature. (D) Confocal view of FITC-dextran-labeled vessels in a
villus positioned at the junction of the middle and distal thirds of the small
intestine of a normal, conventionally raised P14 mouse. (Bars, 25 ?m.)
Villus capillary networks in conventionally raised mice. (A–C) Sections
www.pnas.org?cgi?doi?10.1073?pnas.202604299Stappenbeck et al.
images, taken at 5-?m intervals, a rotating three-dimensional
‘‘movie’’ was constructed for viewing the microvascular network
from multiple perspectives (Fig. 1C).
Vascular network development was first characterized in
conventionally raised mice (i.e., mice with a microbiota). We
focused on the distal half of the intestine, where the density of
greater than in the proximal half (25, 26). Three time points were
selected: postnatal day 7 (P7), P14, and P28. During the suckling
period (P1–P14), the crypt stem cell hierarchy is established (27,
28), the preweaning microbiota is predominated by facultative
anaerobes (26), and mature Paneth cells have yet to emerge (24).
At P7 and P14, the villus microvascular pattern consists of a
simple arch with few or no cross-linking branches (Fig. 1D; n ?
3 mice per time point). The suckling–weaning transition and
crypt–villus morphogenesis are both completed by P28, at which
time the villus capillary network has evolved to a complex
latticework of branched interconnecting vessels (Fig. 1C).
The interval between P14 and P28 corresponds to the tran-
sition between the preweaning microbiota and assembly of a
climax microbial community predominated by obligate anaer-
obes (26). In addition, a full census of differentiated Paneth cells
appears at the base of now fully formed crypts. Paneth cells are
able to sense, respond to, and subsequently shape the intestinal
microbiota through their production and export of antimicrobial
peptides (6). Based on these considerations, we proceeded to
test the contributions of the microbiota and Paneth cells to
The Adult Mouse Germ-Free Small Intestine Has an Arrested Angio-
genic Program That Can Be Rapidly Restarted and Completed After
Bacterial Colonization. Recent DNA microarray-based profiling of
small intestinal gene expression in adult NMRI mice raised
without indigenous microbes (germ-free) suggested that a state
of functional immaturity persists through adulthood when a
microbiota is absent (ref. 29; see Table 1, which is published as
supporting information on the PNAS web site, or http://
gordonlab.wustl.edu/vasculature). Therefore, we compared the
structures of villus capillary networks in 6-week-old male NMRI
germ-free mice, and in age- and gender-matched ex-germ-free
animals colonized for 10 or 30 days with an unfractionated distal
small intestinal microbiota harvested from age- and gender-
matched, conventionally raised adult mice. ‘‘Conventionaliza-
tion’’ resulted in a marked increase in the density of the capillary
network, as evidenced by a statistically significant 2-fold re-
duction in the average number of Syto61-labeled nuclei per
‘‘window’’ (P ? 0.05; Fig. 2 A, B, and D). Microbial induction of
angiogenesis was completed over a short time interval (10 days):
colonization for 30 days produced no statistically significant
changes in network density compared with a 10-day colonization
[Fig. 2D; note that the density of epithelial nuclei per unit of
villus surface area does not change during colonization (data not
the upper third of small intestinal villi. Whole mounts are from the junction between the middle and distal thirds of the small intestines of 6-week-old NMRI
mice (capillaries, green; nuclei, red). (A) Germ-free (GF) mouse. (B) Age-matched ex-germ-free conventionalized (CONV) mouse killed 10 days after colonization
(B. theta) alone. To view three-dimensional rotating images of the capillary networks shown in A–C, see Movies 3–5, which are published as supporting
information on the PNAS web site, or go to http://gordonlab.wustl.edu/vasculature. (D) Quantitation of villus capillary network density. Mean values ? SEM are
(Bars in A–C, 25 ?m.)
Rapid microbial induction of angiogenesis in small intestinal villi of adult ex-germ-free mice. (A–C) Confocal scans of the capillary network present in
Stappenbeck et al.
November 26, 2002 ?
vol. 99 ?
no. 24 ?
B. thetaiotaomicron is a genetically manipulatable, abundant
component of the normal human and mouse distal gut microbial
society (30, 31). It plays an important role in breaking down
otherwise undigestable dietary polysaccharides (9–16, 32). A
10-day colonization of adult germ-free NMRI mice with this
Gram-negative anaerobe recapitulates many of the changes in
gene expression prompted by colonization with an unfraction-
ated ‘‘conventional’’ distal small intestinal microbiota (ref. 29;
see Table 1 or http://gordonlab.wustl.edu?vasculature). It does
so without detectable binding to the epithelium (33). B. thetaio-
taomicron colonization of 6-week-old germ-free NMRI mice for
just 10 days is sufficient to induce a statistically significant (P ?
0.005) 2-fold increase in the density of the villus capillary
network. The resulting decrease in window size was indistin-
guishable from that produced by colonization with an unfrac-
tionated distal small intestinal microbiota (Fig. 2 B–D).
Paneth Cells Produce Factors That Play a Key Role in the Development
of the Villus Microvasculature. We next assessed the effects of
Paneth cells, and of B. thetaiotaomicron, on development of the
capillary network by using a pedigree of germ-free FVB/N
transgenic mice that lacks this cell type. Lineage ablation was
achieved by expressing an attenuated diphtheria toxin A frag-
ment (tox176) under the control of nucleotides ?6,500 to ?34 of
the Paneth-cell-specific mouse cryptdin-2 (CR2) gene (23).
tox176 expression reduced the average Paneth cell census per
crypt by ?95% (data not shown). Electron microscopic studies
disclosed that undifferentiated columnar cells (34) replace Pan-
eth cells. Loss of Paneth cells did not affect cell division rates in
the crypt, as judged by the number S-phase cells/5-?m-thick
sections prepared from normal and transgenic littermates
treated with 5-bromo-2?deoxyuridine (BrdUrd) 90 min before
sacrifice (100 sectioned crypts scored per mouse; n ? 3 mice per
villus architecture, or on the differentiation programs of the
other three small intestinal epithelial cell lineages, as defined by
histochemical stains and immunohistochemical surveys with a
panel of previously characterized (35) antibodies and lectins
(n ? 3 mice studied per group).
The capillary networks of germ-free P28 normal mice and
Paneth cell-deficient CR2-tox176 littermates were compared.
Without Paneth cells, the network was significantly less complex
(2-fold larger average window size in distal small intestinal villi;
n ? 3 mice per group; P ? 0.005; Fig. 3 A, B, and E). This finding
indicates that even in the absence of a microbiota, Paneth cells
produce factors that play a key role in the development of the
B. thetaiotaomicron Stimulates Angiogenesis in the Developing Small
Intestine via Paneth Cells. We then colonized weaning P21 male
germ-free normal and CR2-tox176 mice for 7d with B. thetaio-
taomicron so that animals could be killed at the conclusion of
crypt–villus morphogenesis. Comparisons of the resulting P28
germ-free and B. thetaiotaomicron-mono-associated CR2-tox176
mice revealed that in the absence of Paneth cells, the bacterium
was unable to exert an effect on the density of the distal small
intestinal villus microvasculature (Fig. 3 A, C, and E). In
contrast, in the presence of Paneth cells a 7-day colonization
produced a statistically significant 25% increase in capillary
network density (P ? 0.005; Fig. 3 B, D, and E). The effects of
Paneth cells and B. thetaiotaomicron were additive, resulting in
a ?2-fold difference (P ? 0.005) in network density between
of villi. (A) Germ-free, Paneth cell-deficient P28 male CR2-tox176
mouse. (B) Age- and gender-matched, germ-free, Paneth-cell-
examined 7 days after colonization with B. thetaiotaomicron. (D) P28
nontransgenic mouse killed 7 days after mono-association with
B. thetaiotaomicron. To view three-dimensional rotating images of
the capillary networks shown in A–D, see Movies 6–9, which are
published as supporting information on the PNAS web site, or go to
http://gordonlab.wustl.edu/vasculature. (E) Quantitation of capillary
network density. Mean values ? SEM for each condition are plotted.
The statistical significance of differences between various groups is
noted (Student’s t test). (Bars in A–D, 25 ?m.)
Paneth cell and microbial regulation of angiogenesis. (A–D)
www.pnas.org?cgi?doi?10.1073?pnas.202604299Stappenbeck et al.
colonized normal and CR2-tox176 littermates (Fig. 3 C, D,
This study indicates that indigenous microbes act as environ-
mental agents that shape development of the intestinal villus
microvasculature through Paneth cells. By showing that indige-
nous microbes play a key role in the postnatal development of
host niches they occupy, our findings illustrate of the importance
of the coevolution of animals and their microbial partners,
as well as the concept of ‘‘ecological developmental biology’’
The notion of indigenous gut bacteria contributing to vascular
development is appealing from the perspective of what consti-
tutes the underpinnings of symbiotic host–microbial relation-
ships. Microbes facilitate luminal breakdown of dietary macro-
molecules (to the benefit of both partners). By stimulating
expression of host genes involved in the uptake of the resulting
digestion products (9–16, 32), and by increasing the intestine’s
absorptive capacity through promotion of angiogenesis, the
microbiota provides benefit to the host. Microbial regulation of
angiogenesis also allows the marked increase in bacterial density
and species complexity that occurs during the postnatal period
to be coordinated with an accompanying increase in crypt–villus
units. The net result of such coordination is creation of an
intestinal ‘‘bioreactor’’ capable of fulfilling the growing nutrient
and energetic needs of both partners.
The elegance of this regulatory system is emphasized by the
fact that a key cellular component of the innate immune system
(Paneth cells) is strategically positioned to coordinate develop-
ment of both the microbiota and the microvasculature. The
appearance of Paneth cells coincides with initial colonization of
the gut. Their subsequent differentiation is influenced by the
microbiota, while at the same time their secreted antimicrobial
peptides/proteins effects microbial ecology (6, 24). Moreover,
their placement at the crypt base may allow them to further
influence gut development by affecting the properties of the
stem cell niche.
Paneth expressed factor(s) may act directly on the microvas-
culature, or indirectly by influencing expression of regulators
produced by other epithelial or mesenchymal cell populations.
DNA microarray analysis of laser-capture microdissected epi-
thelial cells harvested from the crypt bases of P28 germ-free
normal and CR2-tox176 mice yielded a list of 63 genes whose
work). No known regulators of angiogenesis are included in this
list, although one gene encodes a newly described (36), micro-
bially regulated (29) member of the angiogenin family of
RNases. Further dissection of the microbial, cellular, and mo-
lecular components of this angiogenesis pathway may provide
important new therapeutic principles for preventing or minimiz-
ing ischemic insults to the gut, facilitating villus regeneration
after injury, improving absorptive function, and/or treating
neoplastic processes. The gnotobiotic normal and transgenic
mice described in this report provide a starting point for this
pathway dissection in a simplified, genetically and environmen-
tally defined system.
We thank David O’Donnell, Maria Karlsson, and Sabrina Wagoner for
superb technical assistance. We are grateful to Richard Hotchkiss and
Mitchell Grayson for suggestions concerning labeling of capillary net-
works. This work was funded in part by grants from the National
Institutes of Health (DK30292 and DK52574) and AstraZeneca. T.S.S.
is the recipient of a National Institutes of Health K08 Career Develop-
ment Grant. L.V.H. is supported by a Burroughs Wellcome Foundation
Career Award in the Biomedical Sciences.
1. Wong, M. H., Saam, J. R., Stappenbeck, T. S., Rexer, C. H. & Gordon, J. I.
(2000) Proc. Natl. Acad. Sci. USA 97, 12601–12606.
2. Booth, C. & Potten, C. S. (2000) J. Clin. Invest. 105, 1493–1499.
3. Cheng, H. (1974) Am. J. Anat. 141, 521–536.
4. Bjerknes, M. & Cheng, H. (1981) Am. J. Anat. 160, 51–63.
5. Ouellette, A. J. & Selsted, M. E. (1996) FASEB J. 10, 1280–1289.
6. Ayabe, T., Satchell, D. P., Wilson, C. L., Parks, W. C., Selsted, M. E. &
Ouellette, A. J. (2000) Nat. Immunol. 1, 113–118.
7. Vaughan, E. E., Schut, F., Heilig, H. G., Zoetendal, E. G., de Vos, W. M. &
Akkermans, A. D. (2000) Curr. Issues Intest. Microbiol. 1, 1–12.
8. Hashimoto, H., Ishikawa, H. & Kusakabe, M. (1998) Anat. Rec. 250, 488–492.
9. Salyers, A. A., Vercellotti, J. R., West, S. E. & Wilkins, T. D. (1977) Appl.
Environ. Microbiol. 33, 319–322.
10. Salyers, A. A., West, S. E., Vercellotti, J. R. & Wilkins, T. D. (1977) Appl.
Environ. Microbiol. 34, 529–533.
11. Salyers, A. A., Harris, C. J. & Wilkins, T. D. (1978) Can. J. Microbiol. 24,
12. Salyers, A. A., Gherardini, F. & O’Brien, M. (1981) Appl. Environ. Microbiol.
13. D’Elia, J. N. & Salyers, A. A. (1996) J. Bacteriol. 178, 7173–7179.
14. D’Elia, J. N. & Salyers, A. A. (1996) J. Bacteriol. 178, 7180–7186.
15. Reeves, A. R., Wang, G. R. & Salyers, A. A. (1997) J. Bacteriol. 179, 643–649.
16. Cho, K. H. & Salyers, A. A. (2001) J. Bacteriol. 183, 7198–7205.
17. Schottelius, M. (1902) Arch. Hyg. Bakteriol. 42, 48–70.
18. Steinert, M., Hentschel, U. & Hacker, J. (2000) Naturwissenschaften 87, 1–11.
19. Hooper, L. V. & Gordon, J. I. (2001) Science 292, 1115–1118.
20. Gilbert, S. F. (2001) Dev. Biol. 233, 1–12.
21. McFall-Ngai, M. J. (2002) Dev. Biol. 242, 1–14.
22. Hooper, L. V., Mills, J. C., Roth, K. A., Stappenbeck, T. S., Wong, M. H. &
Gordon, J. I. (2002) Methods Microbiol. 31, 559–589.
23. Garabedian, E. M., Roberts, L. J. J., McNevin, M. S. & Gordon, J. I. (1997)
J. Biol. Chem. 272, 23729–23740.
24. Bry, L., Falk, P., Huttner, K., Ouellette, A., Midvedt, T. & Gordon, J. I. (1994)
Proc. Natl. Acad. Sci. USA 91, 10335–10339.
25. Ushijima, T., Takahashi, M., Tatewaki, K. & Ozaki, Y. (1983) Microbiol.
Immunol. 27, 985–993.
26. Savage, D. C. (1977) Annu. Rev. Microbiol. 31, 107–133.
27. Schmidt, G. H., Winton, D. J. & Ponder, B. A. (1988) Development (Cambridge,
U.K.) 103, 785–790.
29. Hooper, L. V., Wong, M. H., Thelin, A., Hansson, L., Falk, P. G. & Gordon,
J. I. (2001) Science 291, 881–884.
30. Moore, W. E. C. & Holdeman, L. V. (1974) Appl. Microbiol. 27, 961–979.
31. Salyers, A. A., Bonheyo, G. & Shoemaker, N. B. (2000) Methods 20, 35–46.
32. Hooper, L. V., Midtvedt, T. & Gordon, J. I. (2002) Annu. Rev. Nutr. 22,
33. Bry, L., Falk, P. G., Midtvedt, T. & Gordon, J. I. (1996) Science 273, 1380–1383.
34. Cheng, H. & Leblond, C. P. (1974) Am. J. Anat. 141, 461–480.
35. Hermiston, M. L. & Gordon, J. I. (1995) J. Cell Biol. 129, 489–506.
36. Holloway, D. E., Hares, M. C., Shapiro, R., Subramanian, V. & Acharya, K. R.
(2001) Protein Expression Purif. 22, 307–317.
Stappenbeck et al.
November 26, 2002 ?
vol. 99 ?
no. 24 ?