FGF-2 enhances intestinal stem cell survival and its expression is induced after radiation injury

Courtney W. Houchen, Robert J. George, Mark A. Sturmoski, and Steven M. Cohn

Department of Medicine, Washington University School of Medicine, St. Louis, Missouri 63110


    ABSTRACT
Top
Abstract
Introduction
Materials and methods
Results
Discussion
References

Fibroblast growth factors (FGFs) have mitogenic activity toward a wide variety of cells of mesenchymal, neuronal, and epithelial origin and regulate events in normal embryonic development, angiogenesis, wound repair, and neoplasia. FGF-2 is expressed in many normal adult tissues and can regulate migration and replication of intestinal epithelial cells in culture. However, little is known about the effects of FGF-2 on intestinal epithelial stem cells during either normal epithelial renewal or regeneration of a functional epithelium after injury. In this study, we investigated the expression of FGF-2 in the mouse small intestine after irradiation and determined the effect of exogenous FGF-2 on crypt stem cell survival after radiation injury. Expression of FGF-2 mRNA and protein began to increase at 12 h after gamma -irradiation, and peak levels were observed from 48 to 120 h after irradiation. At all times after irradiation, the higher molecular mass isoform (~24 kDa) of FGF-2 was the predominant form expressed in the small intestine. Immunohistochemical analysis of FGF-2 expression after radiation injury demonstrated that FGF-2 was predominantly found in the mesenchyme surrounding regenerating crypts. Exogenous recombinant human FGF-2 (rhFGF-2) markedly enhanced crypt stem cell survival when given before irradiation. We conclude that expression of FGF-2 is induced by radiation injury and that rhFGF-2 can enhance crypt stem cell survival after subsequent injury.

ionizing radiation; basic fibroblast growth factors; injury repair


    INTRODUCTION
Top
Abstract
Introduction
Materials and methods
Results
Discussion
References

HOMEOSTASIS OF THE NORMAL adult intestinal epithelium is maintained by continuous and rapid replacement of differentiated epithelial cells by replication of undifferentiated epithelial cells (transit cells) located within the crypts of Lieberkühn and subsequent differentiation of their progeny during migration away from the zone of replication (19, 36). However, relatively little is known about the molecular mechanisms that regulate the dynamics of stem cell replication or stem cell fate in the adult intestinal epithelium during either normal epithelial renewal or regeneration of a functional epithelium after injury.

The four principal differentiated cell lineages of the intestinal epithelium are derived from a common multipotent stem cell(s) located near the base of each intestinal or colonic crypt (4). These crypt stem cells divide rarely in the normal intestine to produce a daughter stem cell (self renewal) as well as a more rapidly replicating transit cell. Transit cells, in turn, undergo four to six rapid cell divisions in the proliferative zone located in the lower half of each crypt. Enterocytes, goblet cells, and enteroendocrine cells undergo terminal differentiation as they migrate upward from the zone of proliferation onto the villus epithelium in the small intestine or onto the surface epithelial cuff in the colon (4, 19, 36). Paneth cells differentiate as they migrate to the base of the crypt in the small intestine.

Restoration of normal epithelial architecture and function after intestinal injury induced by a variety of noxious agents (chemical, infectious, radiation, and inflammatory) is a multistep process that must ultimately involve changes in the dynamics of the epithelial stem cell population (34). First, migration of adjacent epithelial cells over the wound reestablishes continuity of the epithelium. Stem cells subsequently proliferate to increase their numbers and to give rise to the more rapidly proliferating transit cell population. The transit cell population then expands rapidly to form a regenerative crypt. A single surviving clonogenic crypt stem cell is sufficient to give rise to a regenerating crypt. If the injury has completely destroyed some crypts, the surviving crypt stem cells can divide to replete the number of viable crypts (34). Finally, normal epithelial function is reestablished by migration of epithelial cells along with the surrounding mesenchymal components to reform normal villus architecture and patterns of epithelial differentiation.

A number of studies (12, 22, 33, 36) have suggested that prostaglandins, peptide growth factors, including members of the fibroblast growth factor (FGF) family, and other regulatory peptides can affect cell survival, proliferation, differentiation, and migration of intestinal epithelial cells in vitro. However, the role of these peptide growth factors and other mediators in regulating the dynamics of crypt epithelial stem cell proliferation has been difficult to study directly because at present there are no specific histological or immunologic characteristics that distinguish stem cells from other crypt epithelial cells. Thus knowledge of the biological characteristics of intestinal stem cells has been largely acquired by inference from experiments using chimeric and transgenic mice (19, 23, 43, 44) and from study of epithelial regeneration after injury directed at replicating cell populations in the gut (34, 42). The response of the crypt stem cell to injury induced by a variety of genotoxic and cytotoxic agents has been primarily studied using a microcolony formation assay based on the capacity of surviving stem cells (clonogens) to regenerate cryptlike foci of replicating epithelial cells that can be scored histologically 3-4 days after injury (34, 45). Intestinal injury induced by ionizing radiation has been the most extensively characterized model. The transit cells within the crypt cease replication after exposure to ionizing radiation. However, these cells continue to migrate out of the crypt and onto the villus. Thus, in the absence of a surviving crypt stem cell, the crypt will disappear. If one (or more) clonogenic stem cell survives irradiation, it will proliferate, ultimately giving rise to an entire regenerative crypt.

FGFs are a family of at least nine heparin-binding polypeptides that have mitogenic activity toward a variety of cell types of mesenchymal, neuronal, and epithelial origin. In addition to their mitogenic activity, these factors have been shown to regulate cellular differentiation in vitro and are involved in neoplasia, angiogenesis, and regulation of morphogenic events in normal embryonic and fetal development (1). FGF-2 is expressed in normal adult tissues as well and may regulate cellular events involved in wound repair (11, 12). A number of studies (6, 15, 16, 20, 21) have shown that FGF-2 enhances survival of a variety of endothelial, epithelial, and hematopoetic cell lines after injury induced by ionizing radiation. The biological effects of FGFs are mediated by binding to and activating cell surface receptor tyrosine kinases encoded by a family of four related genes, FGFR-1, -2, -3, and -4 (32). The ligand-binding specificity of each of these receptors is determined by sequences present in Ig-like domains II and III in the extracellular portion of the receptor molecule. Additionally, multiple receptor isoforms of FGFR-1, -2, and -3 with differing ligand-binding specificities can be expressed as a result of alternative mRNA splicing events that alter the sequence encoding the carboxy-terminal region of Ig-like domain III of each of these receptors. Binding of FGFs to their receptors induces receptor dimerization, transphosphorylation of their cytoplasmic domains, and activation of downstream signaling cascades.

In this study, we examine the expression of FGF-2 in the mouse small intestine after irradiation and the response of the epithelial stem cell population to FGF-2 during radiation injury repair, using the microcolony assay. We report that 1) FGF-2 mRNA and protein levels are elevated in the small intestine after radiation injury, 2) FGF-2 is localized to the mesenchyme surrounding regenerative crypts in irradiated mice, and 3) exogenous recombinant FGF-2 enhances crypt stem cell survival when administered before irradiation but has no effect on epithelial stem cell survival when given only after radiation injury.


    MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and methods
Results
Discussion
References

Animals. FVB/N female mice (Taconic; Germantown, NY) were maintained on a 12:12-h light-dark cycle and fed standard laboratory mouse chow ad libitum. Mice were irradiated at age 8-12 wk in a Gamacel 40 cesium irradiator at 0.96 cGy/min. Some animals received recombinant human FGF-2 (rhFGF-2; 1-4 µg/g body wt ip; gift of Dr. Judith Abraham, Scios-Nova, Mountainview, CA) and/or heparin sulfate (1-4 µg/g body wt; Sigma-Aldrich, St. Louis, MO) at various times either before or after gamma -irradiation. Animals were killed at various times after irradiation and rapidly dissected, as previously described (9). Some mice received 120 mg/kg 5-bromo-2'-deoxyuridine (BrdU) (Sigma-Aldrich) and 12 mg/kg 5-fluoro-2'-deoxyuridine (Sigma-Aldrich) 2 h before death to permit identification of replicating S phase cells by immunohistochemistry. The proximal jejunum was fixed in Bouin's solution and divided into 5-µm segments before paraffin embedding and immunohistochemical analysis. The distal jejunum was snap frozen in liquid nitrogen, and total cellular RNA was prepared for RNase protection analysis from the frozen tissue using TRIzol (GIBCO BRL, Bethesda, MD), according to the manufacturer's suggested protocol.

Measurement of FGF-2 mRNA levels. Jejunal RNA was prepared as described above. Samples (20 µg) of total RNA were hybridized with a 32P-labeled antisense RNA probe corresponding to mouse FGF-2 (24). To quantitate FGF-2 mRNA expression, we also hybridized the 32P-labeled antisense FGF-2 RNA probe to samples containing known amounts of in vitro-transcribed FGF-2 sense-strand RNA. Using the RPA II RNase protection kit (Ambion, Austin, TX) according to the manufacturer's suggested protocol, we hybridized the FGF-2 probe to a 479-nt fragment encompassing the entire coding region of the FGF-2 mRNA. The RNA hybrids were digested with RNase A and T1 at 37°C. The protected RNA fragments were separated by electrophoresis in 7.5 M urea-8% acrylamide sequencing gels. To control for differences in preparation and loading of individual samples, we also performed RNase protection analysis on duplicate samples using a mouse glyceraldehyde 3-phosphate dehydrogenase antisense probe. Gels were dried, and autoradiography was performed using Kodak BioMax MR film.

Western blot analysis. Intestines from treated and control mice were rapidly dissected and frozen in liquid nitrogen. Tissues were then homogenized with a Tekmar Tissuemizer in 1 ml of proteinase inhibitor cocktail (25 µg/ml antipain, 25 µg/ml aprotinin, 25 µg/ml leupeptin, 25 µg/ml chymostatin, 50 µM o-phenanthroline, 10 µg/ml pepstatin A, and 23 mM dithiothreitol in 20 mM TES, pH 7.4). Homogenates were centrifuged at 4°C at 800 g, and the supernatant was frozen on dry ice. The protein concentration of the homogenates was determined using the Bradford method (Bio-Rad, Richmond, CA). One half volume of 3× Laemmli SDS sample buffer was added to samples (final sample buffer concn, 2% SDS, 100 mM dithiothreitol, and 60 mM Tris, pH 6.8) before PAGE. Typically, 50 µg of protein were resolved on 15% polyacrylamide gels containing 0.1% SDS and transferred to polyvinylidene difluoride membrane by semidry blotting, according to the manufacturer's instructions (Bio-Rad). Membranes were blocked with a 5% nonfat dry milk solution in Tris-buffered saline (TBS), pH 7.6, containing 0.1% Tween 20. FGF-2 protein bands were detected by enhanced chemiluminescence (Amersham, Arlington Heights, IL), using a primary antibody to FGF-2 (Ab-2, Oncogene Research Products, Cambridge, MA) and anti-rabbit-horseradish peroxidase secondary antibody (Amersham).

Immunohistochemical techniques. For immunohistochemical localization of mouse FGF-2, we incubated deparaffinized sections of Bouin's fixed tissue with a 1:2,000 to 1:8,000 dilution of a polyclonal rabbit anti-rat FGF-2 prepared against amino acids 1-23 of rat FGF-2 (Chemicon International, Temecula, CA). The specificity of this antibody and its use for immunohistochemistry have been previously described (17, 18). After quenching of endogenous peroxidase activity with 3% hydrogen peroxide, we washed sections with TBS. Bound rabbit anti-FGF-2 was detected by fluorescein-conjugated tyramide signal amplification (TSA direct, Dupont NEN Life Science Products, Boston, MA), according to the manufacturer's suggested protocol, after incubation with biotin-labeled donkey anti-rabbit IgG (Jackson ImmunoResearch Laboratories, West Grove, PA) and subsequent incubation with streptavidin-horseradish peroxidase. To identify replicating cells, we detected BrdU incorporated into S phase cells in deparaffinized sections, using a goat anti-BrdU antibody as previously described (7, 8). Bound anti-BrdU was subsequently visualized with either gold-labeled rabbit anti-goat IgG with silver enhancement (Amersham) or by fluorescence with Texas red-labeled donkey anti-goat IgG (Jackson Immunoresearch Laboratories).

Crypt survival. Crypt survival was measured in young adult (8- to 12-wk-old) mice killed 3.5 days after irradiation, using a modification of the microcolony assay (8). Each mouse received 120 mg/kg BrdU (Sigma-Aldrich) and 12 mg/kg 5-fluoro-2'-deoxyuridine (Sigma-Aldrich) 2 h before death to label the S phase cells. Five-micrometer paraffin sections were prepared from proximal jejunum oriented so that the sections were cut perpendicular to the long axis of the small intestine. For purposes of the microcolony assay, a regenerative crypt was determined to have survived irradiation based on its histological appearance. The viability of each surviving crypt was confirmed by immunohistochemical detection of BrdU incorporation into five or more epithelial cells within each regenerative crypt. A minimum of 12 complete cross sections were scored for each mouse. Because the size of the regenerating crypt may not be the same for each treatment group, the number of surviving crypts per cross section was corrected for crypt size to control for the effect of treatment on the probability of observing a regenerative crypt within a section (37, 38). The width of 15 representative crypts for each animal was measured in longitudinal sections of proximal jejunum at the widest point in each crypt, and the mean surviving crypts per circumference was corrected for the variation in crypt size, as previously described by Potten et al. (38).


    RESULTS
Top
Abstract
Introduction
Materials and methods
Results
Discussion
References

Levels of basic FGF mRNA and protein are increased after radiation injury. The effect of radiation injury on the expression of FGF-2 mRNA in normal adult mice was examined by RNase protection analysis at various times after gamma -irradiation (Fig. 1). In the proximal jejunum of unirradiated mice, FGF-2 mRNA was detectable at low levels (~8 fg/µg total cellular RNA). Levels of FGF-2 mRNA began to increase at 12 h after gamma -irradiation with 13 Gy and rose progressively, reaching peak levels at 72 through 96 h after irradiation that were ~11-fold higher (87 fg/µg total RNA) than observed in unirradiated control mice (Fig. 1). FGF-2 mRNA levels subsequently fell from 96 to 144 h after irradiation to the low levels that were present in unirradiated control mouse small intestine.


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Fig. 1.   Time course of fibroblast growth factor-2 (FGF-2) mRNA expression after irradiation in mouse small intestine. A: FVB/N mice received 13-Gy gamma -irradiation in a cesium irradiator at 0.96 cGy/min. Total cellular RNA was prepared from distal jejunum of mice killed at indicated time points after irradiation. Small intestinal FGF-2 and glyceraldehyde 3-phosphate dehydrogenase (GAPDH) mRNA levels were determined by RNase protection assays. Protected bands were separated by denaturing PAGE, and gels were analyzed by autoradiography. Autoradiographs for FGF-2 were exposed for 7 days and GAPDH for 14 h. B: quantitation of RNase protection assays for FGF-2 shown in A. The intensity of the major protected band for FGF-2 mRNA was determined by scanning densitometry on autoradiographs of RNase protection assays at each time point and compared with the intensity of bands produced with known amounts of in vitro-transcribed sense-strand control mRNA for murine FGF-2. Data were normalized for abundance of GAPDH mRNA present in each sample. Data are average FGF-2 mRNA levels determined from analysis of 2 mice for each time point.

The effect of radiation dose on the expression of FGF-2 mRNA was also determined in the proximal jejunum of mice 72 h after gamma -irradiation, a time point at which regenerating crypts first become apparent (Fig. 2). At doses of 8 Gy or less there were no significant differences in FGF-2 mRNA levels compared with unirradiated control mice. However, at doses of gamma -irradiation above 8 Gy, FGF-2 mRNA levels increased markedly with increasing doses. At 14 Gy, levels of FGF-2 mRNA were ~12-fold higher than present in the proximal jejunum of unirradiated mice.


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Fig. 2.   Radiation dose response of FGF-2 mRNA in mouse small intestine. A: radiation dose response for expression of FGF-2 and GAPDH mRNA was determined by RNase protection assays in mouse small intestine 72 h after irradiation. Protected bands were separated by denaturing PAGE, and autoradiography of gels was performed as described in Fig. 1 legend. B: quantitation of RNase protection assays for FGF-2 in A is shown. Levels of FGF-2 mRNA were determined at each radiation dose as described in Fig. 1B. Data are expressed as average FGF-2 mRNA levels determined from analysis of 2 mice for each radiation dose.

Several isoforms of FGF-2 with different apparent molecular masses have been previously reported (31, 40, 41). The higher molecular mass isoforms of FGF-2 are expressed in many different cell lines in vitro (40). Furthermore, there appears to be tissue-specific differential expression of FGF-2 isoforms in vivo (5). To examine expression of the various FGF-2 protein isoforms in the mouse proximal jejunum after radiation injury, we performed Western blot analysis with anti-FGF-2 directed at an epitope common to all of the reported FGF-2 isoforms (Fig. 3). Low levels of the ~24-kDa isoform were observed in unirradiated control mice. Levels of the ~24-kDa FGF-2 isoform progressively increased after gamma -irradiation (13 Gy), reaching a maximum at 96 h after irradiation. Expression of the ~24-kDa isoform subsequently fell to an intermediate level by 168 h after irradiation. A second fainter ~21.5-kDa band was also observed on Western blots from 72 to 120 h after 13-Gy gamma -irradiation. The 18-kDa FGF-2 isoform resulting from initiation at the AUG codon was not detectable in the proximal jejunum at any time point after irradiation.


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Fig. 3.   Western blot analysis of FGF-2 protein expression in small intestine after gamma -irradiation. Small intestinal lysates were prepared from nonirradiated FVB/N mice and from irradiated mice at times indicated after irradiation with a single dose of 13 Gy. Equal amounts of protein from nonirradiated and irradiated mouse intestinal lysates were separated by electrophoresis on 15% SDS-polyacrylamide gels. Proteins were transferred to polyvinylidene difluoride membranes, and bands corresponding to FGF-2 were detected by enhanced chemiluminescence using rabbit antibodies against FGF-2.

Localization of FGF-2 within small intestine after radiation injury. Immunohistochemical analysis of FGF-2 in the proximal jejunum of unirradiated mice demonstrated scant immunoreactive FGF-2 associated with the basement membrane underlying the crypt and lower portion of the villus epithelium, a pattern similar to that previously found in the developing rat intestine (Fig. 4A) (18). After 13-Gy gamma -irradiation, abundant FGF-2 was observed throughout the lamina propria underlying the lower villus epithelium and beneath the base of the villi (Fig. 4, B, C, E, and G). FGF-2 was also present in the lamina propria beneath the villus base in regions in which regenerating crypts were not present. FGF-2 was not observed in crypt epithelial cells or in the lamina propria associated with the middle or upper villus epithelium. At higher magnification, the immunoreactive FGF-2 appeared to be distributed in a fibrilar pattern, associated with the extracellular matrix surrounding mesenchymal cells within the lamina propria and adjacent to the basilar surface of epithelial cells within regenerating crypts (Fig. 4C). Dual localization of FGF-2 and BrdU confirmed that immunoreactive FGF-2 was predominantly located in the lamina propria surrounding viable actively regenerating crypts containing numerous S phase cells (Fig. 4, E-G). Staining was undetectable in an adjacent control section in which normal preimmune rabbit serum was substituted for the rabbit anti-FGF-2 antibody (Fig. 4D). After irradiation, abundant immunoreactive FGF-2 was also seen surrounding small blood vessels in the submucosa (data not shown).


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Fig. 4.   Immunohistochemical localization of FGF-2 in small intestine of uninjured control and gamma -irradiated mice. Cellular localization of FGF-2 was analyzed in control (A) and gamma -irradiated (13 Gy) (B-G) FVB/N mice. All mice received 5-bromo-2'-deoxyuridine (BrdU) 2 h before death to label S phase cells. Irradiated mice were killed 84 h after irradiation. Sections were incubated with IgG fraction of rabbit anti-rat FGF-2 at a 1:2,000 dilution (A, B, and C) or preimmune rabbit IgG (D) and bound antibody was visualized using biotin-conjugated tyramide amplification technique with 3,3'-diaminobenzidine as chromogenic substrate (brown pigment). In normal mice (A), scant FGF-2 immunoreactivity is present in the basement membrane immediately adjacent to basolateral surface of crypt epithelial cells. In gamma -irradiated mice (B and C), abundant immunoreactive FGF-2 is seen within lamina propria beneath the base of the villi both surrounding regenerating crypts (arrows) and in regions in which regenerating crypts were absent (arrowhead). At higher magnification (C) immunoreactive FGF-2 is seen in a fibrilar pattern consistent with an association with extracellular matrix components in lamina propria. FGF-2 was not detectable within epithelial cells in regenerating crypt. No staining was observed when preimmune rabbit IgG was substituted for rabbit anti-rat FGF-2 (D). E-G: the relationship of FGF-2 to replicating cells within regenerating crypts within the same section is shown. Bound anti-FGF-2 was visualized using fluorescein-conjugated tyramide signal amplification technique (E and G; green fluorescence). S phase cells incorporating BrdU were detected by goat anti-BrdU, and bound antibody was visualized by immunofluorescence using Texas red-labeled donkey anti-goat IgG (F and G) (red fluorescence). E: FGF-2 surrounding a regenerating crypt. F: distribution of S phase cells in same regenerative crypt. G is a dual exposure of E and F and demonstrates that immunoreactive FGF-2 is not present within replicating epithelial cells in regenerating crypt. Bars = 100 µm.

Recombinant FGF-2 enhances intestinal crypt stem cell survival after irradiation. In unirradiated mice, exogenous rhFGF-2 (4 µg/g) given 25 h before analysis had no effect on the number of crypts per cross section or on the fraction of crypt epithelial cells in S phase (data not shown). To determine whether FGF-2 might regulate crypt stem cell survival in response to injury, we treated adult FVB/N mice with rhFGF-2 either before or after gamma -irradiation and crypt stem cell survival was measured using the microcolony assay (Figs. 5 and 6) (8). When FGF-2 was administered intraperitoneally before irradiation, crypt survival increased with increasing duration of FGF-2 administration (Fig. 5). Maximal enhancement of crypt survival was observed when treatment with recombinant FGF-2 was begun 25 h before irradiation. A similar enhancement of crypt survival was observed when mice received either a single dose of rhFGF-2 (4 µg/g) 25 h before irradiation or were given rhFGF-2 (1 µg/g) every 8 h beginning 25 h before irradiation (Fig. 5B). Heparin sulfate, which was coadministered with FGF-2, had no effect on crypt survival when given alone. rhFGF-2 (4 µg/g) did not affect crypt survival when it was given daily beginning immediately after irradiation through the time of death for the microcolony assay (Fig. 6). As has been previously reported by Potten et al. (37, 38), the size of regenerating crypts after gamma -irradiation was significantly larger than the size of crypts in unirradiated control mice. Treatment with rhFGF-2, either 25 h before 13.8-Gy gamma -irradiation or daily beginning immediately postirradiation, did not significantly alter the size of regenerating crypts compared with mice receiving only 13.8-Gy gamma -irradiation (Table 1).


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Fig. 5.   Effects of pretreatment with FGF-2 on crypt survival after radiation injury. A: FVB/N (8-wk-old) mice received a single dose of gamma -irradiation (13.3 Gy). FGF-2-treated mice received a dose of 1 µg/g recombinant human FGF-2 and heparin sulfate (1 µg/g) every 8 h beginning at time indicated before irradiation and continuing until animals were gamma -irradiated. Control group received heparin sulfate (l µg/g) only every 8 h beginning 25 h before irradiation. Each mouse received an injection of BrdU 82 h after irradiation. Animals were killed 2 h later. No. of surviving crypts per jejunal cross section is shown for each dosing schedule. Three mice were analyzed per treatment group. Data are expressed as means ± SE of no. of surviving crypts per cross section. * P < 0.05 compared with control; ** P < 0.005 compared with control. B: FGF-2-treated mice received either recombinant human FGF-2 (rhFGF-2) (1 µg/g) and heparin sulfate (1 µg/g) every 8 h (q8) beginning 25 h before irradiation or a single dose of 4 µg/g rhFGF-2 in heparin sulfate (4 µg/g) at 25 h before irradiation. Heparin-treated control mice received heparin sulfate (1 µg/g) only every 8 h beginning 25 h before irradiation or heparin sulfate (4 µg/g) at 25 h before irradiation, respectively; 13 Gy control mice received sterile saline. No. of surviving crypts per jejunal cross section is shown for each dosing schedule. Three mice were analyzed per treatment group. Data are expressed as means ± SE of the mean no. of surviving crypts per cross section. * P < 0.05 compared with heparin.


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Fig. 6.   Effect of rhFGF-2 administration after irradiation on crypt survival. FVB/N mice (12 wk old) received a single dose of gamma -irradiation (13.8 Gy). FGF-2 treated mice received either rhFGF-2 (4 µg/g) and heparin sulfate (4 µg/g) 25 h before irradiation or every 24 h until death beginning 1 h postirradiation. Control mice received heparin sulfate (4 µg/g) only. No. of surviving crypts per jejunal cross section is shown for each dosing schedule. Twelve complete cross sections were scored for each mouse. Five mice were analyzed per treatment group. Data are expressed as means ± SE of surviving crypts per cross section. * P < 0.005 compared with heparin control. ** Not significant compared with heparin control.

                              
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Table 1.   Effect of irradiation and FGF treatment schedule on crypt dimensions in proximal jejunum

The effect of radiation dose on crypt stem cell survival in the jejunum was investigated in mice treated with FGF-2 (4 µg/g) 25 h before irradiation and in mice treated with the heparin sulfate vehicle alone (Fig. 7). At doses of 10 Gy or less, pretreatment with FGF-2 did not significantly affect crypt survival. However, at doses above 10 Gy, crypt survival decreased with increasing radiation dose. The number of surviving crypts per jejunal cross section at 16 Gy in the heparin sulfate control mice was ~1% of the number of crypts present in jejunal cross sections of unirradiated control mice. Pretreatment with FGF-2 resulted in a marked and progressive enhancement of crypt survival at all doses above 10 Gy. Crypt survival in FGF-2-treated mice was 2.8-fold higher at 12 Gy, 5.3-fold higher at 14 Gy, and 11-fold higher at 16 Gy than observed in mice treated with the heparin sulfate vehicle alone.


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Fig. 7.   Effect of radiation dose on crypt survival in control and rhFGF-2-treated mice. Effect of radiation dose on fractional crypt survival in control mice or mice treated with rhFGF-2 is shown. Adult FVB/N mice (8 wk old) were killed, and crypt survival was scored 31/2 days after a single dose of radiation. FGF-2-treated mice received 4 µg/g rhFGF-2 and heparin sulfate (4 µg/g) at 25 h before irradiation. Control mice received heparin sulfate only, the vehicle for rhFGF-2. Twelve cross sections per mouse from 3 mice were scored for crypt survival for each data point. Data are normalized to no. of BrdU-labeled crypts in unirradiated animals. Data for fractional crypt survival are presented as means ± SE.


    DISCUSSION
Top
Abstract
Introduction
Materials and methods
Results
Discussion
References

In this study we show that radiation injury results in increased expression of FGF-2 mRNA and protein. Levels of FGF-2 mRNA and protein began to rise at 12 h postirradiation when crypt stem cells may be recovering from any cell cycle arrest and are beginning the process of epithelial regeneration. Peak levels of FGF-2 expression occurred from 72 to 96 h after irradiation, a time period when the appearance of regenerative crypts and disruption of epithelial integrity are the predominant morphological features of the reparative process. Increased FGF-2 expression was dependent on the extent of radiation injury. Increased levels of FGF-2 mRNA were only observed at doses above 8 Gy that produce moderate to severe injury to the gastrointestinal epithelium, resulting in progressive loss of viable regenerative crypts and compromise of the epithelial barrier (compare Figs. 2 and 7). Lower doses of radiation injury that suppress replication in the transit cell population but do not impair crypt survival or compromise epithelial barrier function had no effect on levels of FGF-2 mRNA or protein.

The marked increase in FGF-2 mRNA and protein levels that we observed in the small intestine taken together with our immunohistochemical findings suggest that ionizing radiation induces FGF-2 gene expression locally within mesenchymal cells in the lamina propria of the intestine. It is unlikely that FGF-2 is expressed in regenerating crypt epithelial cells or that these epithelial cells induce local production of FGF-2 in the surrounding mesenchyme since FGF-2 was not detectable within epithelial cells after irradiation and because abundant FGF-2 was present both in regions containing regenerating crypts and in areas where regenerating crypts were not present (Fig. 4B). Much of the immunoreactive FGF-2 appeared to be associated with extracellular matrix beneath the base of the villi rather than localized to a particular cell type within the lamina propria. These data are consistent with the known affinity of FGF-2 for heparin sulfate like proteoglycans present in the extracellular matrix. The juxtacryptal localization of FGF-2 suggests that FGF-2 may either act directly on epithelial cells within regenerating crypts by binding to receptors located on the basolateral surface of these epithelial cells or through indirect effects on cells in the mesenchymal tissue that are immediately adjacent to regenerative crypts. It is clear that mesenchymal cells, such as pericryptal fibroblasts, are important in regulating such epithelial functions as crypt cell replication, epithelial migration, and regional differentiation events (19).

The increased expression of FGF-2 in the small intestine induced by irradiation could be due to a direct effect of radiation injury on the transcription of the FGF-2 gene, to a series of sequential events occurring within the cells that express FGF-2, or to a response of FGF-2-expressing cells to cytokines, growth factors, or other signaling molecules elaborated by other cell types after irradiation. Ionizing radiation has been shown to induce expression of FGF-2 directly in several cell lines, including some endothelial and epithelial cell lines in vitro (21, 28). Ionizing radiation and other agents that result in cellular DNA damage can also induce the expression of a number of transcription factors in mammalian cells, such as c-jun, c-fos, EGR-1, and nuclear factor-kappa B (2, 3, 10, 14, 28, 29). However, the induction of these transcription factors occurs within the first few hours after irradiation and is transient, falling back to baseline levels before the time period (48-120 h) when increased expression of FGF-2 was most pronounced (see Fig. 1). Thus it is unlikely that any of these transcription factors are the proximate mediators of the increase in FGF-2 expression that we observe after irradiation in vivo. Alternatively, the radiation-induced increase in FGF-2 expression may result from a cascade of cellular events occurring within FGF-2-expressing cells that is initiated by radiation injury. Another possibility is that the increase in FGF-2 expression after irradiation results from effects of a cytokine or other mediator produced by another cell type on FGF-2-expressing cells in the intestine. For example, expression of cyclooxygenase-1 (COX-1) and production of PGE2 are induced by radiation injury with kinetics (8) that are remarkably similar to the time course of FGF-2 expression after irradiation that we observed in this study. Several other growth factors and cytokines, including tumor necrosis factor-alpha (TNF-alpha ), transforming growth factor-beta 1 (TGF-beta 1), and interleukin-1beta (IL-1beta ), are also produced by endothelial cells, myofibroblasts, and other cell types in response to radiation injury, suggesting the possibility that FGF-2 expression might be regulated through such extracellular signaling mechanisms (21, 46).

The predominant isoform of FGF-2 that we observe after irradiation has an apparent molecular mass of ~24 kDa (Fig. 3). Several investigators (31, 40) have observed that multiple higher molecular mass isoforms of FGF-2 can result from utilization of alternative translation initiation sites at in-frame CUG codons within the FGF-2 mRNA located 5' to the AUG, which is the initiation site for the ~18-kDa isoform. NH2-terminally extended forms of FGF-2 have been isolated from colonic mucosa as well as from other cell types and are mitogenic for intestinal epithelial cell lines in culture (12, 31). Experiments using transgenic mice expressing a single human FGF-2 mRNA suggest that tissue-specific expression patterns of the different molecular mass FGF-2 isoforms are regulated at the translational level (5). The higher molecular mass isoforms of FGF-2 contain a nuclear translocation signal that results in nuclear accumulation of these higher molecular mass isoforms in cultured cells (41). The 18-kDa isoform of FGF-2 lacks this sequence and is primarily found in the cytosol of cells in culture. We find that after irradiation much of the immunoreactive FGF-2 is associated with the extracellular matrix (Fig. 4). Nuclear localization of immunoreactive FGF-2 was not observed. These data taken together with the results of the Western blot analysis showing that only the higher molecular mass ~24- and ~21.5-kDa isoforms are detectable in the mouse small intestine (Fig. 3) suggest that, in contrast to cells in culture, the high-molecular-mass isoforms of FGF-2 are synthesized and released from cells in response to radiation injury in vivo. In experiments using transfected cell lines that express either the 18-kDa isoform or the high-molecular-mass isoform of FGF-2 exclusively, the different isoforms of FGF-2 had distinct biological effects on motility, integrin expression, serum growth requirements, and FGF receptor expression in addition to their shared biological activities (13, 26). The high-molecular-mass isoforms of FGF-2 are thought to mediate at least some of their effects on cellular function independent of activation of cell surface receptor tyrosine kinases. Thus it is possible that the expression of endogenous 24-kDa FGF-2 after irradiation can affect the response of stem cells to radiation injury through mechanisms that are different from the effects that we observe on crypt stem cell survival with the exogenously administered rhFGF-2.

It is likely that exogenous rhFGF-2 alters the sensitivity of epithelial stem cells to subsequent radiation-induced damage since rhFGF-2 maximally enhanced crypt stem cell survival when it was given 25 h before irradiation. A number of mechanisms could account for this radioprotective effect of rhFGF-2 on intestinal epithelial stem cells. 1) FGF-2 might reduce radiation-induced damage by increasing cellular levels of substances that prevent formation or increase breakdown of reactive oxygen intermediates that are thought to mediate damage of DNA and other macromolecules within the cell. However, studies (15) on cell lines in culture have found no effect of FGF-2 on the spectrum or amount of DNA damage produced by gamma -irradiation. 2) FGF-2 could directly enhance injury-repair mechanisms in epithelial stem cells. 3) FGF-2 could cause cell-cycle arrest, allowing time for repair to occur in otherwise replicating cell populations. Ionizing radiation can induce transient arrest of the cell cycle at checkpoints in G1 and/or G2 in a variety of intestinal and nonintestinal cell lines (6, 27). Such cell cycle checkpoints have been shown to play an important role in the repair of DNA damage and enhancement of survival of cells exposed to genotoxic agents (6, 27). Several studies (6, 15, 16) have shown that treatment of cell lines with exogenous FGF-2 and transfection of cell lines with plasmids expressing FGF-2 enhance cell survival and repair of DNA damage after irradiation. Furthermore, radioresistance induced by the high-molecular-mass isoforms of FGF-2 was associated with hypophosphorylation of the cell cycle regulatory protein p34 cdc2 and prolongation of G2 in HeLa cells (6). 4) FGF-2 might enhance crypt survival by protecting stem cells from undergoing programmed cell death after irradiation. Apoptosis is thought to be the primary mechanism for regulating the number of intestinal epithelial stem cells in the crypt during normal epithelial renewal and to be the predominant biological response of crypt epithelial cells to low levels of damage that occur chronically in the small intestine and colon (39). However, this may not be an appropriate biological response in epithelial stem cells when injury is more severe and intestinal barrier function is compromised.

Although expression of FGF-2 is markedly increased after irradiation, exogenous rhFGF-2 had no effect on crypt stem cell survival when administration was begun immediately after radiation injury. It is possible that FGF-2 is not able to prevent stem cell death when radiation injury has already occurred or that the effects of endogenously produced FGF-2 on crypt stem cells during epithelial regeneration may already be maximal. Another possibility is that the pharmacological effects of exogenously administered rhFGF-2 may be different from the effects of endogenous FGF-2 present in the gut after injury.

A variety of cytokines and other intracellular regulatory molecules, including TGF-beta 3, KGF, TNF-alpha , IL-1, IL-11, and the prostaglandin derivatives dimethyl-PGE2 and misoprostol, when given before injury, have been shown to protect epithelial stem cells within the intestine from cell death induced by radiation or other cytotoxic agents (22, 25, 30, 35, 37). Here, we show that exogenous rhFGF-2 given before irradiation also enhances clonogenic stem cell survival in response to subsequent injury. It is interesting that the expression of COX-1 and the production of intestinal PGE2 are increased after radiation injury (8) with a time course that is nearly identical to the kinetics of endogenous FGF-2 expression that we report in this study. Furthermore, inhibition of PGE-2 synthesis through COX-1 resulted in decreased stem cell survival after gamma -irradiation. Additionally, the radiation dose response of FGF-2 expression was remarkably similar to the dose-dependent increase for COX-1 and PGE2 after irradiation that we previously reported (8). Increased expression of FGF-2, COX-1, and PGE2 were only observed at radiation doses that cause a significant decrease in the number of regenerating crypts within the intestinal epithelium and result in significant damage to or loss of the epithelial barrier. Thus increased expression of endogenous FGF-2 appears to be a component of a coordinated response to radiation injury in the intestine.

A variety of biological functions have been suggested for FGF-2 in injury repair, including stimulation of angiogenesis, enhanced remodeling of the extracellular matrix, regulation of fibroblast and smooth muscle proliferation, and stimulation of epithelial migration (11, 12). Although the biological consequences of increased endogenous FGF-2 expression in the small intestine after radiation injury have not yet been determined, our data raise the possibility that, in addition to its other activities in injury repair, a potential function of FGF-2 may be to enhance survival of intestinal epithelial stem cells after genotoxic or cytotoxic damage.


    ACKNOWLEDGEMENTS

This study was supported by National Institutes of Health Grants R01 DK-50924 and R01 HD-31914 (both to S. M. Cohn) and by a generous gift from the Ladies Auxiliary of the Veterans of Foreign War, Missouri Chapter. S. M. Cohn was also supported by a Glaxo Institute of Digestive Health Basic Research Award and an American Gastroenterological Association Procter and Gamble Research Scholar Award. C. W. Houchen is a recipient of a Robert Wood Johnson Minority Medical Faculty Development Award.


    FOOTNOTES

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Address for reprint requests: S. M. Cohn, Division of Gastroenterology and Hepatology, Univ. of Virginia Health Sciences Center, PO Box 10013, Charlottesville, VA 22906.

Received 27 January 1998; accepted in final form 25 September 1998.


    REFERENCES
Top
Abstract
Introduction
Materials and methods
Results
Discussion
References

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