The inflammatory bowel diseases (IBDs) Crohn’s disease and ulcerative colitis are the most common chronic gastrointestinal (GI) illness in children and adolescents Approximately, 1.4 million Americans have IBD, and more than 30,000 new cases are diagnosed in the United States annually. IBD is a key risk factor for bone loss and fractures in children1–7 and adults.8–13 Multiple factors are thought to contribute to IBD effects on bone, including nutritional status,3,14–19 reduced physical activity/muscle,1,20 glucocorticoid use,1 and inflammatory cytokines.21,22
The skeleton is a highly dynamic tissue that is regulated by local, systemic, and environmental cues that modify osteoblast (bone formation) and/or osteoclast (bone resorption) activities, and IBD impacts bone regulation at all levels: environmentally through intestinal barrier breaks and/or altered gut microbial composition, systemically through circulation of gut immune cells and cytokines throughout the body, and locally by causing inflammation of extra-intestinal organs (such as the bone marrow). Chronic inflammatory diseases are, in general, associated with increased risk for osteoporosis.23 Treatment of these diseases with steroids can further contribute to bone loss and fracture risk in IBD8,24,25; however, low bone density is evident in men recently diagnosed with IBD before steroid use,26 and young adults with IBD exhibit little or no correlation between steroid use and bone density status. Many studies demonstrate that IBD is associated with increased production of pro-inflammatory factors, such as tumor necrosis factor–alpha (TNF-α), interleukin (IL)-1β, IL-17, and IL-6. Treatment of colitis patients with infliximab, a TNF-α–inactivating antibody, can be successful in treating colitis and improves bone mineral density (spine and femur) in patients with Crohn’s disease.21,27–29 Most studies do not address changes in bone architecture, lineage selection, gene expression, or local cytokine expression.
Multiple mouse strains with targeted immune alterations, such IL-2−/− 30 and IL-10−/−,31 develop colitis and thus are used as models of IBD. Intestinal luminal microbes are needed for the development of colitis in these genetic models.30,32,33 Enterohepatic Helicobacter species exist in the intestinal tract and livers of humans, other mammals, and birds.34–36 It was subsequently found that Helicobacter hepaticus infection of mice with altered immune function could trigger IBD. Re-derivation of these strains to Helicobacter-free status will reduce or eliminate the development of colitis, whereas challenge with H. hepaticus triggers the development of colitis.37–40 Two days after infection, H. hepaticus is a minor component of the microbiota, but after 14 days, it becomes the dominant member of the microbial community.41 Helicobacter hepaticus infection induces Th1 and Th17 immune responses marked by elevated interferon gamma (IFN-γ) and IL-17 levels, respectively.42
IBD affects both men and women,1 although the magnitude of bone loss is suggested to differ by gender.43 Specifically, a cross-sectional study of 51 patients with IBD found that gender (being male) was negatively associated with spine and femur T-scores in ulcerative colitis patients,43 suggesting that the extent of disease-induced bone loss was related to gender. In the present study, we examined the effect of H. hepaticus–induced colitis in IL-10−/− mice on GI and bone health to determine if there is a gender difference in bone loss and disease severity. Our findings indicate that male mice display greater bone loss and disease severity compared with female mice infected with H. hepaticus and suggest different pathological and inflammatory responses to colitis in this model.
MATERIALS AND METHODS
To examine the effect of experimental IBD on bone health, 14-week-old IL-10−/− male and female littermate mice were infected with H. hepaticus and examined 6 weeks later. Age-matched noninfected littermates (controls) were sham-infected with sterile tryptic soy broth. To examine the influence of estrogen deficiency, IL-10−/− mice were ovariectomized at 12 weeks of age and gavaged with H. hepaticus at 14 weeks and examined 6 weeks later. Adult mice, at the plateau of their growth rate, were used to reduce the contribution of confounding effects on bone growth; also, at this age, bone length has stabilized, allowing comparative analyses among animals. Breeding pairs of Helicobacter-free C57BL/6 IL-10−/− were housed in a specific pathogen-free environment and given autoclaved food, bedding, and water. Cage changes were performed in a laminar flow hood. Experimental mice were transferred to the University Research Containment Facility at Michigan State University at 12 weeks and housed under the same conditions as the breeding pairs. Animals were housed up to 4 animals per cage in specific pathogen free conditions and were negative (by PCR analysis) for Helicobacter before the experiment (data not shown). Mice were maintained on a 12:12-h light–dark cycle at 23°C and had food and water ad libitum. The Michigan State University Institutional Animal Care and Use Committee approved all animal procedures.
Helicobacter hepaticus was obtained from American Type Culture Collection (ATCC 51449; Manassas, VA). Helicobacter hepaticus was maintained on tryptic soy agar supplemented with 5% sheep blood (HemoStat Laboratory, Dixon, CA). Helicobacter hepaticus cultures were maintained at 37°C in a microaerobic environment generated in vented GasPak jars without catalyst that were evacuated to −20 mm Hg and equilibrated with a gas mixture consisting of 80% N2, 10% CO2, and 10% H2.
Murine Infection with Helicobacter hepaticus
Cultured H. hepaticus was harvested from agar plates and resuspended in a small volume of sterile tryptic soy broth. The inoculum optical density (at 600 nm wave length) was measured and diluted with sterile tryptic soy broth to an optical density of 1.0 to 2.0. Male and female littermate mice at 14 weeks of age were inoculated with a single dose of 0.3-mL bacterial suspension. Bacteria were introduced directly to the stomach with a 24-guage ball-tipped gavage needle. Control mice were inoculated with sterile tryptic soy broth.
Detection of Helicobacter hepaticus in Mouse Feces
Colonization was confirmed 1 week after infection. DNA was isolated from fecal pellets using DNeasy Kit (Qiagen, Valencia, CA). Single-stage PCR amplification was performed with primers (B38) 5′-GCATTTGAAACTGTTACTCTG-3′ and (B39) 5′-CTGTTTTCAAGCTCCCC-3′, which produce a 417-bp amplicon used to detect H. hepaticus.44 PCR analysis was performed using 5-μL DNA, 20 pmol of each primer, and iQ SYBR Green Supermix (Bio-Rad, Hercules, CA). Cycling condition were 30 cycles of 30 seconds at 95°C, 45 seconds at 54°C, and 45 seconds at 72°C. PCR products were visualized by agarose gel electrophoresis.
Cecum Histology and Scoring
To monitor and confirm colitis induction, a scoring system was developed based on previous reports45–47 using live mouse measures and cecum histology. Disease status in live mice evaluated fecal pellet consistency (0–10, ranging from normal to loose to bloody diarrhea), appearance of rectal bleeding, and weight loss. For histomorphometric evaluation, paraffin-embedded cecum sections were stained with hematoxylin–eosin and imaged at ×4, ×10, and ×25 magnification. At high magnification, 3 areas were imaged, coded, and analyzed/scored blinded to the condition and gender of the mouse. Active disease was evaluated by measures of crypt distortion, numbers of lymphoid aggregates, numbers of lymphoid cells and neutrophils, and mucosal inflammation/ulceration severity.
Crypt distortion measurements were obtained at the ileocecocolic junction and were scored as follows: 0, straight crypts; 2, crypts with slight distortion; 4, distorted crypts with mild hyperchromasia with fewer goblet cells; 6, larger areas of pronounced crypt distortion, more pronounced hyperchromasia with very few goblet cells; 8, crypt drop-out, marked hyperchromasia and no goblet cells. In each section, the number of lymphoid aggregates were counted and the abundance of lymphoid cells was scored as follows: 0, 0 to 10 lymphoid cells counted; 2, 11 to 30 cells; 4, 31 to 100; and 6, more than 100 lymphoid cells, respectively, and counted within the submucosal layer of the cecum. Neutrophils were scored as follows: 0, no neutrophils present; 5, few neutrophils; and 10, numerous neutrophils in section. Disease severity was examined using the entire cecum region and was scored as follows: 0, normal; 2, mild, small focal areas (1–5 crypts) of disease/inflammation limited to the lamina propria; 4, moderate, multifocal with areas of disease/inflammation extended into the submucosa; 6, severe with ulcers covering large areas of mucosa less than 20 crypts wide; 8, very severe with ulcers covering large areas of mucosa more than 20 crypts; and 10, extremely severe with ulcers covering large areas of mucosa more than 20 crypts and signs of bacteria or fungal invasion observed. Total overall disease score was obtained by adding the live mouse scores with the values obtained from the histological analyses. To further confirm colitis, RNA was extracted from the proximal colon to assess cytokine expression levels as previously described48 and noted below. Similarly, tissue was processed for cytokine protein levels.
Helicobacter infection was detected histologically on silver-stained sections stained by the Steiner method. Sections were scored as number of positive crypts per visual field (>100 crypts counted per mouse cecum sample), 4 photographs per cecal section. The percentage of infected crypts was a scoring value. The number of visualized Helicobacter organisms per crypt was also counted and used as a score, and this ranged from 1 to 5. In the cases of heavily infected male cecum crypts, the score of 10 was given to each crypt that had too numerous bacteria to count. The total Steiner stain score was the percentage of infected crypts added to the score of Helicobacter organisms.
Enzyme-linked Immunosorbent Assay Assay
To determine the tissue cytokine levels, colons were snap frozen in liquid nitrogen and homogenized in phosphate-buffered solution. TNF-α, IFN-γ, and IL-17A levels in homogenates were analyzed using ELISA kits from eBioscience, Inc (San Diego, CA). Cytokine levels were normalized to the total protein and expressed as picogram per milligram of total protein as described before.49
Immediately after euthanasia, tibias were cleaned of muscle and connective tissue, snap frozen in liquid nitrogen and stored at −80°C. Frozen tibias were crushed under liquid nitrogen conditions with a Bessman Tissue Pulverizer (Spectrum Laboratories, Rancho Dominguez, CA). RNA was isolated using TriReagent (Molecular Research Center, Cincinnati, OH), and integrity was assessed by formaldehyde–agarose gel electrophoresis. Complementary DNA was synthesized by reverse transcription using Superscript II Reverse Transcriptase Kit and oligo dT(12-18) primers (Invitrogen, Carlsbad, CA) and amplified by real-time PCR with iQ SYBR Green Supermix (Bio-Rad), and gene-specific primers were synthesized by Integrated DNA Technologies (Coralville, IA). Hypoxanthine guanine phosphoribosyl transferase mRNA levels that do not fluctuate with disease were used as an internal control. Amplicon specificity was confirmed by melting curve, size, and sequence analysis. Primers for real-time PCR were designed or previously described: Runx2, OC, AP, Trap5b, HPRT,48 ColI (5′-AGAAAGGATCTCCTGGTGCTGAT-3′ and 5′-AAGCCTCTTTCTCCTCTCTGACC-3′), OPG (5′GAAGAAGATCATCCAAGACATTGAC-3′ and 5′-TCCATAAACTGAGTAGCTTCAGGAG-3′), and RankL (5′-TTTGCAGGACTCGACTCTGGAG-3′ and 5′-TCCCTCCTTTCATCAGGTTATGAG-3).
Microcomputed Tomography Bone Imaging
Fixed femurs were scanned using a GE Explore Locus microcomputed tomography system at a voxel resolution of 20 μm obtained from 720 views. Beam angle of increment was 0.5, and beam strength was set at 80 peak kV and 450 μA. Each run consisted of control (broth treated) and H. hepaticus–treated bones, and a calibration phantom to standardize grayscale values and maintain consistency. On the basis of autothreshold and isosurface analyses of multiple bone samples, a fixed threshold (800) was used to separate bone from bone marrow. Trabecular bone analyses were performed in a region of trabecular bone defined at 0.17 mm (∼1% of the total length) distal to the growth plate of the proximal femur extending 2 mm toward the diaphysis, excluding the outer cortical bone. Trabecular bone mineral content, bone volume fraction, thickness, spacing, and number values were computed by a GE Healthcare MicroView software application for visualization and analysis of volumetric image data. Cortical measurements were performed in a 2 × 2 × 2-mm cube centered midway down the length of the bone using a threshold of 1,000 to separate bone from marrow.
Femur Histomorphometry and Dynamic Measures
Femurs were fixed in 10% formalin and transferred to 70% ethanol after 24 hours. Fixed samples were processed on an automated Thermo Electron Excesior tissue processor for dehydration, clearing, and infiltration using a routine overnight processing schedule. Samples were then embedded in Surgipath-embedding paraffin on a Sakura Tissue Tek II-embedding center. Paraffin bocks were sectioned at 5 μm on a Reichert Jung 2030 rotary microtome. Slides were stained for TRAP activity and counterstained with hematoxylin according to manufacturer protocol (387A-1KT; Sigma, St Louis, MO). Osteoblast and osteoclast surface area was measured and expressed as a percentage of total bone surface in the femur trabecular region ranging from the growth plate to 2 mm distal.
For dynamic histomorphometric measures of bone formation, mice were injected intraperitoneally with 200 μL of 10 mg/mL calcein (Sigma) dissolved in sterile saline at 7 and 2 days before harvest. L3–L4 vertebrae were fixed in formalin at the time of harvest and then transferred to 70% ethanol 48 hours later. Vertebrae were then embedded, sectioned, and examined under UV light. Five images were taken, and the distance between the calcein lines (bone formation rate) and their length along the bone surface was measured and used to calculate mineral apposition rate.
All measurements are presented as the mean ± standard error. Statistically significant (α = 0.05) effects of H. hepaticus or gender or H. hepaticus × gender (which would indicate gender altering H. hepaticus effects or vice versa) were determined using factorial (2-way) analysis of variance (ANOVA) with SPSS statistical software (Chicago, IL). Student’s t test (assuming equal variance) using Microsoft Excel (Microsoft, Redmond, WA) was also used to determine significance where noted. For categorical, ordinal data comparisons (such as disease severity, crypt distortion, or Steiner staining scores), 2-tailed Mann–Whitney U tests were used (http://vassarstats.net/index.html) to determine statistical significance at P < 0.05.
Male and female littermate IL-10−/− mice were infected with H. hepaticus at 14 weeks of age and harvested at 20 weeks of age to assess the impact of this model on bone density and architecture at an age where bone remodeling predominates over growth. Both femur and vertebrae were examined because these sites can differ in responses to treatments/disease. As expected, femurs from male mice had more trabecular bone volume fraction and mineral content and density, trabecular number, trabecular thickness, and cortical bone area than that of femurs from female mice (Fig. 1; Table 1). Similar to other models of colitis,48 H. hepaticus infection resulted in a significant decrease in male trabecular bone parameters, such as bone volume fraction (BV/TV −37%) but only slightly decreased most cortical parameters (not reaching statistical significance). The outer cortical perimeter, however, did decrease significantly after infection. Surprisingly, compared with the male bone parameters, none of the female bone parameters (trabecular or cortical) were affected by H. hepaticus infection (Fig. 1; Table 1). Normalization of femur bone volume fraction values by individual mouse weights, to correct for differences in body weight, still showed similar findings (Fig. 1). Two-way ANOVA analysis identified gender and H. hepaticus as having a significant (P < 0.001) effect on femur bone volume fraction and identified a significant effect caused the interaction of gender × H. hepaticus (P < 0.001).
Examination of vertebrae trabecular structure did not identify a difference between male and female control mice (Fig. 2; Table 2). Although vertebrae can display unique responses to treatments/disease, H. hepaticus infection caused somewhat similar changes/trends in vertebrae to that seen in femurs. Specifically, H. hepaticus significantly reduced vertebral trabecular bone volume only in male mice (−46%) but not in female mice (Fig. 2). Two-way ANOVA identified H. hepaticus as having a major effect on vertebral bone volume (P < 0.05) and that the interaction of H. hepaticus and gender displayed a trend toward affecting vertebral bone volume (P = 0.06). Other vertebral trabecular parameters were also altered in male mice but not in female mice (Table 2).
Tibial RNA was used to determine if markers of osteoblast (bone anabolic cells) or osteoclast (bone catabolic cells) differentiation were affected by H. hepaticus. HPRT, which is not modulated by colitis or inflammation, was used as a housekeeping gene. Osteoblast maturation is characterized by their ability to secrete type 1 collagen-rich extracellular matrix during early stages of maturation and by markers of osteoblast lineage/differentiation, such as expression of alkaline phosphatase and osterix. All 3 of these genes were reduced in the bones of male mice treated with H. hepaticus (Fig. 3) but not in the bones of female mice treated with H. hepaticus, consistent with the gender-dependent bone loss after infection. Osteoclast activity markers, Trap5b, and osteoclast regulators, RANKL (activator) and OPG (inhibitor), were also examined. Trap5b levels did not differ among conditions (Fig. 3), consistent with Trap5b serum levels (data not shown). RANKL levels displayed a trend toward decreasing in treated male mice (P = 0.08), and OPG levels were significantly decreased in treated male mice. To further assess the impact of H. hepaticus on osteoblasts and osteoclasts, cell surface area in trabecular bone regions were measured by histomorphometry. Helicobacter hepaticus infection did not cause any significant changes in osteoblast or osteoclast surface area or mineral apposition rate. We did notice that H. hepaticus–treated female mice demonstrated a trend toward increased osteoclast number and decreased mineral apposition rate (Table 2); the consequence of this is not clear as the female mice did not lose bone.
With the supposition that colitis severity could differ between genders and in this way contribute to gender differences in bone pathology, general body and intestinal parameters were examined. As expected, H. hepaticus–infected male mice lost a significant amount of body weight (−5%; Fig. 4). In contrast, infected female mice did not lose weight. Correspondingly, infected male mice lost tibialis muscle mass compared with control male mice (−8%; Fig. 4), whereas female mice did not lose muscle mass. Two-way ANOVA indicated that gender and H. hepaticus infection had a significant effect on body and muscle mass and that a gender × H. hepaticus effect was significant for muscle mass and nearly significant for body mass. As expected, muscle and body mass was strongly impacted (P < 0.001) by gender, with control male mice having more compared with control female mice.
Next, we examined colitis severity at 6 weeks using cecum histology and scoring (Fig. 5) where the total disease score was based on the additive total of crypt/mucosal structure, lymphoid aggregate number, lymphoid cell number, neutrophil presence, ulcer severity scores, and assessment of live animals (see Materials and Methods for scoring specifics). Although control male and female mice had similar scores (7.0 ± 2 versus 7.2 ± 2, respectively), H. hepaticus–treated mice had elevated disease severity scores (Fig. 5). Most interestingly, the total disease score for H. hepaticus–treated male mice was nearly 2-fold greater than treated female mice (38.9 ± 6.00 versus 21.6 ± 4.49, respectively). Two-way ANOVA indicated a strong gender, H. hepaticus, and gender × H. hepaticus effect on the level of disease severity (all displayed a P < 0.05). Increased disease severity in male versus female mice was evident for all individual parameters, including mucosal structure (6.0 ± 0.6 versus 3.6 ± 0.3, respectively) and lymphoid aggregates (4.5 ± 0.9 versus 2.4 ± 0.6, respectively).
To further assess and quantitate disease severity, colon cytokine levels were measured. Consistent with the cecum disease score, the colon of male mice treated with H. hepaticus displayed a significant increase in cytokine production, whereas the infected female mice did not show elevated cytokine production (Fig. 6). Specifically, levels of TNF-α, produced by several immune and nonimmune cell types, were increased in male but not female mouse colon (152 ± 51 versus 40.0 ± 10, respectively) and were affected by disease and displayed a trend to be influenced by gender and gender × H hepaticus (P < 0.06). Based on previous report that H. hepaticus infection induces a Th1 and Th17 immune response, we also measured interferon gamma (IFN-γ) and IL-17 levels, respectively.42 Consistent with a Th1 and Th17 response, IFN-γ and IL-17A levels were elevated in H. hepaticus–treated male but not female mouse colon. Two-way ANOVA determined that IFN-γ and IL-17 were regulated by gender, H. hepaticus infection, and the combination of both parameters (gender × H. hepaticus).
In our study, male and female mice were infected with the same quantity of H. hepaticus; however, variations in the amount of H. hepaticus retained and thriving within the GI tract could impact disease severity (higher levels being associated with greater disease severity). Examination of H. hepaticus levels in mouse stool 1 week after infection indicated that similar levels of the bacteria were present in both male and female mice (Fig. 7). However, 6 weeks after infection, H. hepaticus DNA levels were greater in the colons of male compared with female mice, suggesting that H. hepaticus did not thrive over time in the female mice (Fig. 7). This was supported by greater Steiner staining of helicobacter bacteria in male versus female colon histology sections. Thus, greater levels of H. hepaticus and disease severity are associated with increase bone loss. We further carried out regression analyses looking at male and female relationships to disease score; results could not be combined/pooled because of large gender differences, including male bone and body parameters that are significantly greater than female mice. Regression analyses and Pearson’s correlation coefficients indicated a significant relationship between disease severity and bone loss (femur BV/TV) (Fig. 8) and weight, fat pad, and muscle loss in male mice but not in female mice.
To determine if loss of sex steroids could make female mice more vulnerable to H. hepaticus infection and bone loss, littermate female mice were ovariectomized and given H. hepaticus. Ovariectomy did not cause cecal inflammation in the IL-10–deficient female mice. Interestingly, infection of ovariectomized mice with H. hepaticus did not increase cecum inflammation. Consistent with the literature, analyses of femoral trabecular bone volume (Fig. 9) and other parameters (data not shown) indicate that ovariectomy induces significant trabecular bone loss, by more than 50%, whereas infection with H. hepaticus did not cause further bone loss.
Our findings demonstrate that H. hepaticus–infected male mice display greater bone loss and cecal inflammation (disease severity) compared with female mice. In clinical studies, IBD is clearly a risk factor for bone loss and fractures in children1–7 and adults,8–13 but the risk factors for this outcome are not completely clear. Our finding that there is a gender component to bone loss is somewhat surprising because there are few clinical reports describing the contribution of gender to the extent of bone loss in IBD but the findings are variable. One study indicates that for patients with ulcerative colitis, but not Crohn’s disease, male gender is negatively correlated with bone density and that male patients have significantly lower lumbar spine and distal femur T-scores than female patients.43 However, another study found no significant differences in gender or age between fracture and nonfracture patients with IBD, but older age, female sex, and osteoporosis are associated with hospitalization for fractures in patients with IBD.50 The variations in findings suggest a complexity in the relationship between gender and IBD induced bone loss.
Analyses of bone markers (histomorphometry and RNA analyses) in the male mice support a reduction in osteoblast activity. Our findings of suppressed osteoblast activity are supported by studies in our laboratory and others, which indicate a reduction in bone formation in other mouse models of IBD.48,51 Suppressed osteoblast activity was also observed in a study conducted in young (8 and 12 week) male IL-10–deficient mice compared with wild-type mice52; the same study found that some of the mice, which were housed under a standard (not pathogen free) environment, developed spontaneous colitis and that this was associated with greater bone loss, consistent with our findings in male IL-10–deficient mice infected with H. hepaticus. Regarding osteoclast activity, we did not observe suppression in osteoclast markers, which would have suggested an overall reduction in bone remodeling as has been reported clinically.53 Other animal studies show variable results with regard to osteoclast activity.51,52 In addition to differences in mouse age among the studies, we found that osteoclast regulators, both activators and inhibitors, were suppressed in the male mice and may possibly negate any directional effect in our study.
Our studies demonstrate in the H hepaticus–infected IL-10–deficient mouse colitis model that disease severity is greater in adult male mice compared with female mice, which correlates with the extent of bone loss. Clinically, there is a relationship between IBD severity and/or duration and bone loss and fractures,13,43,54 which is not surprising, given that IBD can cause weight loss, reduced nutrient intake, and systemic inflammation. A few clinical studies have directly examined the contribution of gender to colitis severity. Some studies do not demonstrate a relationship between gender and disease severity, but note that age is reciprocally related to disease severity.55,56 Others have identified female IBD patients as having a lower remission rate and less immunosuppressive medications than male patients.57 Thus, the contribution of gender to IBD severity is not completely clear and difficult to assess at the mechanistic level clinically because variations in factors, such as diet, genetics, gut microbiota, and sex hormone levels, can confound interpretations.
Mouse models of colitis allow identification of potential mechanisms regulating disease severity and complications, such as bone loss. Helicobacter hepaticus–induced colitis is one model where factors contributing to disease severity are being identified, including signaling factors, such as SMAD3,58 gut permeability, and Th1 inflammatory activity.59 Our studies use H. hepaticus–infected IL-10−/− mice on the C57BL/6 background. Interestingly, IL-10–deficient mice in this background are less prone to develop H. hepaticus–induced colitis compared with IL-10–deficient Balbc or 129 mice, which develop moderate and severe colitis, respectively.42 The mechanisms accounting for the different strain responses to H. hepaticus have yet to be identified but likely involve variations in immune responses. Studies have also shown that certain host genetic loci, such as Cdcs160 and Nod2,59 can affect H. hepaticus virulence/severity.
Consistent with a Th1 and Th17 immune response to H. hepaticus infection, male mice displayed increases in gut IFN-γ and IL-17 levels, respectively.42 This was not seen in female mice. Studies indicate that elevated cytokines are associated with disease severity,42 thus the lack of cytokine induction in female H. hepaticus–infected mice is consistent with the reduced disease severity. Thus, male gender increases the severity of H. hepaticus–induced colitis in C57BL/6 mice. Livingston et al61 also examined the role of gender in H. hepaticus–induced colitis in A/JCr mice, but in contrast to our studies, female A/JCr mice displayed greater severity than male mice. Differences in mouse strain (A/JCr versus C57BL/6), mouse age of infection (3 weeks of age versus 14 weeks of age in our study), and length of inoculation (3 months versus 1.5 months) likely contribute to the different results seen in the 2 studies. This reinforces the role of age and mouse strain on responses to H. hepaticus infection and colitis severity.
How could gender influence disease severity? Our findings indicate that H. hepaticus survival and virulence within the GI tract is gender dependent. Differences in the intestinal environment could contribute to this and include differences in gut immune surveillance, gut acidity, and even bile acid composition. The latter can affect the growth and proteome of H. hepaticus in a way that could alter virulence, inflammation, host adaptation, and colonization location.62 Regression analysis indicates that the disease score, an indicator of intestinal inflammation, is negatively correlated with bone volume in male mice. This was not observed in female mice. One possible cause of this gender difference is that a high/threshold disease score may be necessary to initiate bone loss because the majority of infected male mice displayed a disease score well above female mice (an average of 39 for male versus 21 for female mice).
The lack of visible bone loss in H. hepaticus–infected ovariectomized mice suggests that estrogen may not be required for the protective effects at this age and time point. However, this interpretation should be taken with caution for 2 main reasons. First, it is possible that the significant bone loss in the ovariectomized mice prevents observation of additional reductions (from an already low bone volume). Second, estrogenic effects (immune, neurological, and bile) can be permanent or take time to deplete in adult mice; therefore, the ovariectomized mice could still be benefiting from previous estrogen effects. The lack of bone loss in H. hepaticus–infected ovariectomized mice correlates with the lack of cecal inflammation, consistent with pro-inflammatory cytokines being required to induce IBD in this model.63 Our studies suggest that female mice are protected from gut inflammation or alternatively male mice, with testosterone, are more susceptible to infection. Understanding the role of sex steroids in regulating bone loss caused by H. hepaticus is difficult because interpretations have to made in light of bone loss that already occurs in the absence of estrogen or testosterone. Taken together, the data indicate that long-term infection by H. hepaticus in the GI tract is reduced in female compared with male mice and could contribute to differences in disease severity and ultimately bone loss. The role of sex steroids in this process is complex and a focus of ongoing studies.
The authors thank Jing Zhang for her critical review of the manuscript, and the Investigative Histology Laboratory in the Department of Physiology, Division of Human Pathology at Michigan State University for their assistance with histological analyses.
1. Burnham JM, Shults J, Semeao E, et al.. Whole body BMC in pediatric Crohn disease: independent effects of altered growth, maturation, and body composition. J Bone Miner Res. 2004;19:1961–1968.
2. Sylvester FA, Davis PM, Wyzga N, et al.. Are activated T cells regulators of bone metabolism in children with Crohn disease? J Pediatr. 2006;148:461–466.
3. Semeao EJ, Jawad AF, Zemel BS, et al.. Bone mineral density in children and young adults with Crohn's disease. Inflamm Bowel Dis. 1999;5:161–166.
4. Gokhale R, Favus MJ, Karrison T, et al.. Bone mineral density assessment in children with inflammatory bowel disease. Gastroenterology. 1998;114:902–911.
5. Sylvester FA, Wyzga N, Hyams JS, et al.. Natural history of bone metabolism and bone mineral density in children with inflammatory bowel disease. Inflamm Bowel Dis. 2007;13:42–50.
6. Sylvester FA. IBD and skeletal health: children are not small adults! Inflamm Bowel Dis. 2005;11:1020–1023.
7. Semeao EJ, Stallings VA, Peck SN, et al.. Vertebral compression fractures in pediatric patients with Crohn's disease. Gastroenterology. 1997;112:1710–1713.
8. Bernstein CN, Leslie WD, Leboff MS. AGA technical review on osteoporosis in gastrointestinal diseases. Gastroenterology. 2003;124:795–841.
9. Bartram SA, Peaston RT, Rawlings DJ, et al.. Multifactorial analysis of risk factors for reduced bone mineral density in patients with Crohn's disease. World J Gastroenterol. 2006;12:5680–5686.
10. Bjarnason I, Macpherson A, Mackintosh C, et al.. Reduced bone density in patients with inflammatory bowel disease. Gut. 1997;40:228–233.
11. Pollak RD, Karmeli F, Eliakim R, et al.. Femoral neck osteopenia in patients with inflammatory bowel disease. Am J Gastroenterol. 1998;93:1483–1490.
12. van Hogezand RA, Hamdy NA. Skeletal morbidity in inflammatory bowel disease. Scand J Gastroenterol Suppl. 2006;(243):59–64.
13. van Staa TP, Cooper C, Brusse LS, et al.. Inflammatory bowel disease and the risk of fracture. Gastroenterology. 2003;125:1591–1597.
14. Siffledeen JS, Siminoski K, Steinhart H, et al.. The frequency of vitamin D deficiency in adults with Crohn's disease. Can J Gastroenterol. 2003;17:473–478.
15. Siffledeen JS, Fedorak RN, Siminoski K, et al.. Randomized trial of etidronate plus calcium and vitamin D for treatment of low bone mineral density in Crohn's disease. Clin Gastroenterol Hepatol. 2005;3:122–132.
16. Sentongo TA, Semaeo EJ, Stettler N, et al.. Vitamin D status in children, adolescents, and young adults with Crohn disease. Am J Clin Nutr. 2002;76:1077–1081.
17. Vaisman N, Dotan I, Halack A, et al.. Malabsorption is a major contributor to underweight in Crohn's disease patients in remission. Nutrition. 2006;22:855–859.
18. Pappa HM, Gordon CM, Saslowsky TM, et al.. Vitamin D status in children and young adults with inflammatory bowel disease. Pediatrics. 2006;118:1950–1961.
19. Duggan P, O'Brien M, Kiely M, et al.. Vitamin K status in patients with Crohn's disease and relationship to bone turnover. Am J Gastroenterol. 2004;99:2178–2185.
20. Lee N, Radford-Smith G, Taaffe DR. Bone loss in Crohn's disease: exercise as a potential countermeasure. Inflamm Bowel Dis. 2005;11:1108–1118.
21. Paganelli M, Albanese C, Borrelli O, et al.. Inflammation is the main determinant of low bone mineral density in pediatric inflammatory bowel disease. Inflamm Bowel Dis. 2007;13:416–423.
22. Strober W, Fuss IJ. Proinflammatory cytokines in the pathogenesis of inflammatory bowel diseases. Gastroenterology. 2011;140:1756–1767.
23. Armour KE, Van THRJ, Grabowski PS, et al.. Evidence for a pathogenic role of nitric oxide in inflammation-induced osteoporosis. J Bone Miner Res. 1999;14:2137–2142.
24. Vestergaard P, Krogh K, Rejnmark L, et al.. Fracture risk is increased in Crohn's disease, but not in ulcerative colitis. Gut. 2000;46:176–181.
25. Compston J. Management of glucocorticoid-induced osteoporosis. Nat Rev Rheumatol. 2010;6:82–88.
26. Sakellariou GT, Moschos J, Berberidis C, et al.. Bone density in young males with recently diagnosed inflammatory bowel disease. Joint Bone Spine. 2006;73:725–728.
27. Bernstein M, Irwin S, Greenberg GR. Maintenance infliximab treatment is associated with improved bone mineral density in Crohn's disease. Am J Gastroenterol. 2005;100:2031–2035.
28. Miheller P, Muzes G, Zagoni T, et al.. Infliximab therapy improves the bone metabolism in fistulizing Crohn's disease. Dig Dis. 2006;24:201–206.
29. Pazianas M, Rhim AD, Weinberg AM, et al.. The effect of anti-TNF-alpha therapy on spinal bone mineral density in patients with Crohn's disease. Ann N Y Acad Sci. 2006;1068:543–556.
30. Sadlack B, Merz H, Schorle H, et al.. Ulcerative colitis-like disease in mice with a disrupted interleukin-2 gene. Cell. 1993;75:253–261.
31. Kuhn R, Lohler J, Rennick D, et al.. Interleukin-10-deficient mice develop chronic enterocolitis. Cell. 1993;75:263–274.
32. Morrissey PJ, Charrier K. Induction of wasting disease in SCID mice by the transfer of normal CD4+/CD45RBhi T cells and the regulation of this autoreactivity by CD4+/CD45RBlo T cells. Res Immunol. 1994;145:357–362.
33. Sellon RK, Tonkonogy S, Schultz M, et al.. Resident enteric bacteria are necessary for development of spontaneous colitis and immune system activation in interleukin-10-deficient mice. Infect Immun. 1998;66:5224–5231.
34. Solnick JV, Schauer DB. Emergence of diverse Helicobacter
species in the pathogenesis of gastric and enterohepatic diseases. Clin Microbiol Rev. 2001;14:59–97.
35. On SL, Hynes S, Wadstrom T. Extragastric Helicobacter
species. Helicobacter. 2002;7(Suppl 1):63–67.
36. Fox JG. The non-H. pylori
helicobacters: their expanding role in gastrointestinal and systemic diseases. Gut. 2002;50:273–283.
37. Chin E, Dangler CA, Fox JG, et al.. Helicobacter hepaticus
infection triggers inflammatory bowel disease in T cell receptor alpha/beta mutant mice. Comp Med. 2000;50:586–594.
38. Erdman S, Fox JG, Dangler CA, et al.. Typhlocolitis in NF-kappa B-deficient mice. J Immunol. 2001;166:1443–1447.
39. Cahill RJ, Foltz CJ, Fox JG, et al.. Inflammatory bowel disease: an immunity-mediated condition triggered by bacterial infection with Helicobacter hepaticus
. Infect Immun. 1997;65:3126–3131.
40. Burich A, Hershberg R, Waggie K, et al.. Helicobacter
-induced inflammatory bowel disease in IL-10- and T cell-deficient mice. Am J Physiol Gastrointest Liver Physiol. 2001;281:G764–G778.
41. Kuehl CJ, Wood HD, Marsh TL, et al.. Colonization of the cecal mucosa by Helicobacter hepaticus
impacts the diversity of the indigenous microbiota. Infect Immun. 2005;73:6952–6961.
42. Fox JG, Ge Z, Whary MT, et al.. Helicobacter hepaticus
infection in mice: models for understanding lower bowel inflammation and cancer. Mucosal Immunol. 2011;4:22–30.
43. Ardizzone S, Bollani S, Bettica P, et al.. Altered bone metabolism in inflammatory bowel disease: there is a difference between Crohn's disease and ulcerative colitis. J Intern Med. 2000;247:63–70.
44. Young VB, Knox KA, Pratt JS, et al.. In vitro and in vivo characterization of Helicobacter hepaticus
cytolethal distending toxin mutants. Infect Immun. 2004;72:2521–2527.
45. Bleich A, Mahler M, Most C, et al.. Refined histopathologic scoring system improves power to detect colitis QTL in mice. Mamm Genome. 2004;15:865–871.
46. Pratt JS, Sachen KL, Wood HD, et al.. Modulation of host immune responses by the cytolethal distending toxin of Helicobacter hepaticus
. Infect Immun. 2006;74:4496–4504.
47. Wirtz S, Neufert C, Weigmann B, et al.. Chemically induced mouse models of intestinal inflammation. Nat Protoc. 2007;2:541–546.
48. Harris L, Senagore P, Young VB, et al.. Inflammatory bowel disease causes reversible suppression of osteoblast and chondrocyte function in mice. Am J Physiol Gastrointest Liver Physiol. 2009;296:G1020–G1029.
49. Patial S, Shahi S, Saini Y, et al.. G-protein coupled receptor kinase 5 mediates lipopolysaccharide-induced NFkappaB activation in primary macrophages and modulates inflammation in vivo in mice. J Cell Physiol. 2011;226:1323–1333.
50. Ananthakrishnan AN, McGinley EL, Binion DG, et al.. Fracture-associated hospitalizations in patients with inflammatory bowel disease. Dig Dis Sci. 2011;56:176–182.
51. Hamdani G, Gabet Y, Rachmilewitz D, et al.. Dextran sodium sulfate-induced colitis causes rapid bone loss in mice. Bone. 2008;43:945–950.
52. Dresner-Pollak R, Gelb N, Rachmilewitz D, et al.. Interleukin 10-deficient mice develop osteopenia, decreased bone formation, and mechanical fragility of long bones. Gastroenterology. 2004;127:792–801.
53. Sylvester FA, Wyzga N, Hyams JS, et al.. Natural history of bone metabolism and bone mineral density in children with inflammatory bowel disease. Inflamm Bowel Dis. 2007;13:42–50.
54. Reffitt DM, Meenan J, Sanderson JD, et al.. Bone density improves with disease remission in patients with inflammatory bowel disease. Eur J Gastroenterol Hepatol. 2003;15:1267–1273.
55. Roth LS, Chande N, Ponich T, et al.. Predictors of disease severity in ulcerative colitis patients from Southwestern Ontario. World J Gastroenterol. 2010;16:232–236.
56. Lee GJ, Kappelman MD, Boyle B, et al.. Role of sex in the treatment and clinical outcomes of pediatric patients with inflammatory bowel disease. J Pediatric Gastroenterol Nutr. 2012;5:701–706.
57. Blumenstein I, Herrmann E, Filmann N, et al.. Female patients suffering from inflammatory bowel diseases are treated less frequently with immunosuppressive medication and have a higher disease activity: a subgroup analysis of a large multi-centre, prospective, internet-based study. J Crohns Colitis. 2011;5:203–210.
58. McCaskey SJ, Rondini EA, Clinthorne JF, et al.. Increased presence of effector lymphocytes during Helicobacter hepaticus
-induced colitis. World J Gastroenterol. 2012;18:1459–1469.
59. Biswas A, Liu YJ, Hao L, et al.. Induction and rescue of Nod2-dependent Th1-driven granulomatous inflammation of the ileum. Proc Natl Acad Sci USA. 2010;107:14739–14744.
60. Buchler G, Wos-Oxley ML, Smoczek A, et al.. Strain-specific colitis susceptibility in IL10-deficient mice depends on complex gut microbiota-host interactions. Inflamm Bowel Dis. 2012;18:943–954.
61. Livingston RS, Myles MH, Livingston BA, et al.. Sex influence on chronic intestinal inflammation in Helicobacter hepaticus
-infected A/JCr mice. Comp Med. 2004;54:301–308.
62. Okoli AS, Raftery MJ, Mendz GL. Effects of human and porcine bile on the proteome of Helicobacter hepaticus
. Proteome Sci. 2012;10:27.
63. Kullberg MC, Rothfuchs AG, Jankovic D, et al.. Helicobacter hepaticus
-induced colitis in interleukin-10-deficient mice: cytokine requirements for the induction and maintenance of intestinal inflammation. Infect Immun. 2001;69:4232–4241.
Keywords:© Crohn's & Colitis Foundation of America, Inc.
bone; osteoblast; colitis; gender; intestine; inflammation