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. Author manuscript; available in PMC: 2019 Apr 11.
Published in final edited form as: Cell Host Microbe. 2018 Mar 22;23(4):447–457.e4. doi: 10.1016/j.chom.2018.03.002

Nutritional support from the intestinal microbiota improves hematopoietic reconstitution after bone marrow transplantation in mice

Anna Staffas 1, Marina Burgos da Silva 1, Ann E Slingerland 1, Amina Lazrak 1, Curtis J Bare 2, Corey D Holman 2, Melissa D Docampo 1,3, Yusuke Shono 1, Benjamin Durham 4, Amanda J Pickard 5, Justin R Cross 5, Christoph Stein-Thoeringer 1, Enrico Velardi 1, Jennifer J Tsai 1, Lorenz Jahn 1, Hillary Jay 1, Sophie Lieberman 1, Odette M Smith 1, Eric G Pamer 1,6,7,8, Jonathan U Peled 1,6,9, David E Cohen 2, Robert R Jenq 10, Marcel RM van den Brink 1,6,9,11,*
PMCID: PMC5897172  NIHMSID: NIHMS953922  PMID: 29576480

Summary

Bone marrow transplantation (BMT) offers curative potential for patients with high-risk hematologic malignancies, but the post-transplantation period is characterized by profound immunodeficiency. Recent studies indicate that the intestinal microbiota not only regulates mucosal immunity, but can also contribute to systemic immunity and hematopoiesis. Using antibiotic-mediated microbiota depletion in a syngeneic BMT mouse model, here we describe a role for the intestinal flora in hematopoietic recovery after BMT. Depletion of the intestinal microbiota resulted in impaired recovery of lymphocyte and neutrophil counts, while recovery of the hematopoietic stem- and progenitor-compartments and the erythroid lineage were largely unaffected. Depletion of the intestinal microbiota also reduced dietary energy uptake and visceral fat stores. Caloric supplementation through sucrose in the drinking water improved post-BMT hematopoietic recovery in mice with a depleted intestinal flora. Taken together, we show that the intestinal microbiota contribute to post-BMT hematopoietic reconstitution in mice through improved dietary energy uptake.

Keywords: Hematopoiesis, bone marrow cell transplantation, immune reconstitution, microbiota, nutrition

eTOC blurb

Intestinal bacteria can exert effects on systemic hematopoiesis. Staffas et al show that the intestinal flora contributes to hematopoietic recovery after bone-marrow transplantation (BMT) through improved dietary-energy uptake. The findings suggest possible clinical intervention strategies for improved BMT outcomes.

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Introduction

Allogeneic bone marrow transplantation (BMT) is an important therapy with curative potential for patients with high-risk hematologic malignancies, but the post-transplantation period is characterized by profound immunodeficiency. Delayed immune reconstitution leaves the patient susceptible to opportunistic infections and is an important contributor to transplant-related morbidity and mortality (Maury et al., 2001; Small et al., 1999). In addition, the donor immune system can exert graft-vs-tumor activity and robust reconstitution has been associated with protection against relapse of the underlying malignancy (Goldberg et al., 2017; Ishaqi et al., 2008; Michelis et al., 2014). Impaired recovery of immune function, particularly in the lymphocyte compartment, has also been described after autologous BMT (Guillaume et al., 1998) and is associated with poor overall survival (Porrata and Markovic, 2004). Thus, understanding the mechanisms by which the immune system reconstitutes and how environmental exposures might influence this process is an important objective in improving outcomes after BMT.

The intestinal microbiota plays an important role in shaping both mucosal and systemic immunity (Belkaid and Hand, 2014). Presence of an intestinal microbiota primes myelopoiesis and reduces susceptibility to infection in mice (Balmer et al., 2014; Clarke et al., 2010; Deshmukh et al., 2014; Khosravi et al., 2014; Tada et al., 1996). Polysaccharide A derived from the intestinal commensal B. fragilis increases CD4 T cell counts (Mazmanian et al., 2005) and the microbiota can promote the generation of hematopoietic stem- and progenitor cells (Iwamura et al., 2017; Josefsdottir et al., 2016; Luo et al., 2015). In addition, depletion of the intestinal flora reduces mobilization of hematopoietic stem cells (HSC) in an experimental model of cytokine-induced peripheral HSC mobilization (Velders et al., 2004). It is thus well established that the intestinal bacteria or signals derived from them contribute to hematopoiesis. All of these studies, however, were done under steady-state conditions, and to our knowledge the role of the intestinal microbiota during the critical expansion of the hematopoietic system that occurs following BMT has not been studied.

We have previously reported that the composition of the intestinal microbiota is associated with risk for relapse of malignancy after BMT (Peled et al., 2017) and that low diversity of the post-transplant intestinal flora is associated with increased transplant-related mortality and worse overall survival in BMT patients (Taur et al., 2014). Since both relapse and transplant-related mortality are outcomes that are inversely correlated with a robust immune reconstitution (Goldberg et al., 2017; Ishaqi et al., 2008; Kim et al., 2004; Michelis et al., 2014), we hypothesized that an intact gut microbiota promotes immune reconstitution after BMT. By performing syngeneic transplantation in antibiotic (abx)-treated mice and mice with an intact intestinal flora we demonstrate links between the microbiota, nutrition, and post-transplant hematopoiesis.

Results

Depletion of the intestinal microbiota impairs post-BMT hematopoiesis

To test the role of the intestinal microbiota in immune reconstitution after BMT, we performed syngeneic BMT (B6 → B6) after lethal irradiation in specific pathogen free (SPF) mice with an intact intestinal flora and in mice treated with two different abx regimens: ampicillin + enrofloxacin (AE) and vancomycin + amikacin (VA) administered in drinking water (Fig. 1A). Ampicillin and enrofloxacin are both relatively well absorbed in the intestine (Eriksson and Bolme, 1981; Heinen, 2002) while vancomycin and amikacin both have poor oral bioavailability with negligible systemic effects (Jagannath et al., 1999; Tedesco et al., 1978). Both treatments reduced the colonic bacterial abundance 1000-fold compared to untreated control mice (Fig. 1B). After BMT, we found a dramatic reduction in peripheral white blood cell (WBC) count recovery in mice treated by either of the abx regimens, while platelet- (PLT) and red-blood-cell (RBC) counts were less affected (Fig. 1C). The reduction in total WBC count could be explained to some extent by lower counts of neutrophils and monocytes, but the most dramatic difference was a 3-fold decrease in lymphocytes. Abx-treated mice had lymphocyte counts below the normal range (Fig. 1C) and 5- and 3-fold reductions in B and T cell lineages, respectively (Fig. 1D). Consistent with an impaired hematopoietic recovery, AE- and VA-treated mice also had lower bone marrow cellularity 28 days after BMT compared to untreated mice (Fig. 1E). Importantly, abx treatment also lowered WBC and lymphocyte counts in an allogeneic, minor-MHC-antigen disparate BMT model (129 → B6, Fig. S1A). To assess the functional implication of this lymphopenia we infected mice intravenously with Listeria monocytogenes 21 days after BMT following a 3-day abx washout (Fig. S1B). AE-treated mice had higher bacterial load in the spleen 3 days after infection compared with untreated controls, indicating a functional immune deficit in the mice with a depleted intestinal flora (Fig. S1C).

Fig. 1. Depletion of the intestinal microbiota impairs immune reconstitution after bone marrow transplantation and sensitizes mice to sub-lethal irradiation.

Fig. 1

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(A) Experimental procedure of BMT. PB – Peripheral blood analysis. (B) Quantification of bacterial 16S rRNA in fecal samples from untreated control (n = 10), ampicillin + enrofloxacin (AE)-treated (n = 10), and vancomycin + amikacin (VA)-treated mice (n = 5) 14 days after BMT. NTC = Non Template Control. (C) White blood cells (WBC), red blood cells (RBC), platelets (PLT), lymphocytes (LYMPH), neutrophils (NEUT), and monocytes (MONO), (D) Flow-cytometry analysis of B and T cells in peripheral blood after BMT and (E) Total bone marrow cellularity 28 days after BMT in control (n = 10), AE-treated (n = 8), and VA-treated mice (n = 10). (F) Experimental procedure of semi-lethal irradiation. (G) Survival and (H) Representative images of hematoxylin- and eosin-stained bone-marrow vertebrae sections from untreated mice (left panel) and AE-treated mice (middle panel) 21 days after 750cGy radiation and from an age-matched untreated unirradiated control mouse (right panel). Scale bar 100μm. (I) Experimental procedure of control/resistant fecal microbiota transfer (FMT) and subsequent BMT. (J) Quantification of bacterial 16S rRNA copies in fecal samples from mice at day 0. Control (C) FMT with or without AE-treatment and resistant (R) FMT with or without AE-treatment (n = 5 per group). (K) Total bone marrow cellularity 28 days after BMT and (L) WBC, RBC, PLT, LYMPH, NEUT, and MONO counts after BMT in mice given a control FMT without (n = 10) or with (n = 9) AE-treatment and mice given a resistant FMT without (n = 10) or with (n = 10) AE-treatment. Significance levels are comparison of AE-treated Ctrl-FMT and AE-treated Res-FMT.

Shaded areas in (C) and (L) indicate normal ranges. * P < 0.05, ** P < 0.01, *** P < 0.001, n.s. – Not significant. Results represent at least two independent experiments. Data is presented as mean ± SEM. See also Figure S1.

Depletion of the intestinal microbiota sensitizes mice to semi-lethal irradiation

While survival after a lethal dose of radiation requires transplantation of unexposed donor bone marrow, hematopoietic reconstitution can also be modeled by sub-lethal irradiation and subsequent endogenous hematopoietic recovery without transplant. Depletion of the flora with the AE- or VA-abx regimen sensitized mice to a 750 cGy semi-lethal irradiation dose (Fig. 1F); 60% of untreated mice survived up to 60 days after irradiation while all AE-treated mice and 90% of VA-treated mice died around day 25 after irradiation (Fig. 1G). All abx-treated mice had lower lymphocyte counts and VA-treated mice had lower neutrophil counts when compared to untreated mice 14 days after irradiation (Fig. S1D). The time of death (mean 24 days, range 18 – 31 days) indicated hematopoietic failure (Williams et al., 2010) and moribund mice had an acellular bone marrow (cells from both hind legs totaled 3.3 – 7.3 × 106 which is less than 10% of the normal count) but no signs of infection or sepsis (no ascites or bacteria in peripheral blood or tissues). Furthermore, necropsy of AE-treated mice that were still alive 21 days after irradiation showed centrilobular hepatocellular atrophy and fatty change (Fig. S1E) consistent with hypoxia due to prolonged severe anemia (Fig. S1F). Mice with a depleted flora also showed reduced hematopoietic regeneration compared to untreated mice (Fig. 1H), possibly explaining the reduced survival of mice with a depleted flora after semi-lethal irradiation. Thus, abx-mediated depletion of the intestinal flora impairs both hematopoietic reconstitution after syngeneic BMT and autologous recovery after semi-lethal irradiation.

Impaired post-transplant hematopoiesis in abx-treated mice is mediated by flora depletion

We next assessed whether the detrimental effect of abx treatment on hematopoietic reconstitution is mediated by the intestinal microbiota. The effects observed after oral administration of either absorbed (AE) or non-absorbed (VA) drugs (Fig. 1A – H) suggested a microbiota-mediated effect. To verify that the impaired hematopoiesis was due to depletion of the microbial flora rather than a direct systemic effect of the abx, we utilized a colony of mice that harbor a beta-lactam-resistant microbiota by virtue of having been maintained for years under continuous abx administration (Caballero et al., 2017). One group of mice was given a fecal microbiota transfer (FMT) of the resistant flora (Res-FMT), while control mice were given an FMT with normal flora from SPF mice (Ctrl-FMT). The mice within each group were then subjected to either AE-treatment or no abx and underwent BMT (Fig. 1I). As expected, Res-FMT mice had sustained abundance of fecal bacteria despite AE-treatment (Fig. 1J) and 16S rRNA sequencing showed a diverse flora similar to that of the Res-FMT donor mice (Fig. S1G and H). Plasma concentrations of ampicillin and enrofloxacin were not lower in Res-FMT recipients, demonstrating that the transferred resistant flora was not degrading the abx (Fig. S1I). Bone marrow cellularity, WBC counts, and frequencies of lymphocytes and myeloid cells were effectively rescued in the AE-treated Res-FMT mice compared to AE-treated Ctrl-FMT mice (Fig. 1K and L), demonstrating that the impaired post-BMT hematopoiesis was due to depletion of the intestinal flora and not direct effects of the abx on hematopoiesis.

Depletion of the intestinal flora impairs lymphoid and myeloid differentiation

To further determine the effects of flora depletion on post-transplant hematopoiesis we analyzed the hematopoietic stem- and progenitor-cell compartment 28 days after BMT. Despite an almost 50% decrease in total bone marrow cells, the number of long- and short-term hematopoietic stem cells (LT-HSC and ST-HSC), multi-potent-, common lymphoid-, and common myeloid progenitors (MPP, CLP and CMP) were not consistently reduced in the mice with a depleted flora (AE-, VA-treated, or Ctrl-FMT mice treated with AE) when compared to mice with an intact intestinal flora (all untreated groups and AE-treated Res-FMT mice) (Fig. 2 and Fig. S2). The reduced total bone marrow cellularity could be explained by reduced numbers of both myeloid- and B lymphoid cells, while reductions in T cells were observed as lower number of thymocytes (Fig. 2 and Fig S2). Closer evaluation of differentiating B cells in the bone marrow and spleen revealed lower numbers of all cell subsets, except transitional B cells in the spleen, as well as lower total splenic cellularity in the mice with a depleted flora compared to mice with intact flora (Fig. S3A – D). Similarly, all T cell subsets in the thymus were reduced in abx treated mice compered to untreated mice (Fig. S3E and F). Thus, the effects of flora depletion on post-transplant hematopoiesis present mainly as changes in expansion and differentiation of more mature cells.

Fig. 2. Abx-mediated depletion of the intestinal microbiota predominantly suppresses hematopoietic differentiation.

Fig. 2

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Spider plot (left panel) and heat map (right panel) of number of long-term hematopoietic stem cells (LT-HSC, Lin–ckit+Sca1+CD150+CD48–), short-term hematopoietic stem cells (ST-HSC, Lin–ckit+Sca1+CD150−CD48–), multi-potent progenitors (MPP, Lin–ckit+Sca1+CD150–CD48+), common lymphoid progenitors (CLP, Lin–IL7Rα+ckit+), common myeloid progenitors (CMP, Lin–ckit+Sca1–FcγRlow/–CD34+), megakaryocyte-erythroid progenitors (MEP, Lin–ckit+Sca1–FcγR–CD34–), granulocyte-monocyte progenitors (GMP, Lin− ckit+Sca1–FcγR+CD34+), bone marrow (BM) myeloid cells (CD11b+), total thymocytes, bone marrow (BM) B cells (B220+), and peripheral blood red blood cells (RBC) 28 days after BMT in untreated (n = 10), AE-treated (n = 8), VA-treated (n = 10), Ctrl-FMT mice without (n = 10) or with (n = 9) AE-treatment and Res-FMT mice without (n = 10) or with (n = 10) AE-treatment.

Presented as percentage in relation to untreated ctrl mice (No FMT no Abx). Results represent two independent experiments. * P < 0.05, ** P < 0.01, *** P < 0.001. Data is presented as mean ± SEM. See also Figure S2 and Figure S3.

Morphologic assessment of bone marrow smears revealed that the ratio of myeloid to erythroid cells was decreased in AE- and VA-treated mice compared to untreated BMT recipients (Fig. S3G). Similarly, reductions were observed in the granulocyte-monocyte progenitor (GMP) compartment in AE- and VA-treated mice compared to untreated mice while numbers of megakaryocyte-erythroid progenitors (MEP) and, as previously mentioned, RBC counts were not significantly perturbed in the mice with a depleted flora (Fig. 2 and Fig. S2D). In addition, in vitro colony-forming cultures showed equal frequencies of myeloid colony forming units that together with the reduced total bone marrow cellularity in mice with a depleted flora indicated reduced numbers of GMPs (Fig. S3H).

These results show that flora depletion impaired myeloid and lymphoid differentiation while largely sparing the stem- and progenitor-cell compartments and erythroid lineage (Fig. 2). Thus, the flora is likely to influence expansion and differentiation steps in hematopoiesis.

To assess an alternative hypothesis that initial homing of donor cells to the marrow was impaired by flora depletion we analyzed bone marrow and spleen compositions 16h after injection of cells. Transfer of CFSE-labeled whole-bone-marrow cells or of GFP+ Lineage−Sca-1+ckit+ (LSK) cells showed comparable homing in untreated mice and mice treated for 5 days with AE (Fig. S3I and J), indicating that the gut flora did not contribute to initial homing.

Abx-mediated depletion of the intestinal flora decreases energy harvest from the diet and reduces visceral adipose tissue

In addition to showing impaired hematopoietic recovery, we noticed that the mice treated with abx lost around 20% of their body weight during the first weeks after transplant while untreated mice lost only about 3% (Fig. 3A). Although all treatment groups regained their baseline weight at the end of the 28-day experiment, abx-treated mice had less visceral adipose tissue (VAT) compared to untreated mice as assessed by the amount of periovarian fat (Fig. 3B), which is one of the largest VAT depots in female mice (Chusyd et al., 2016). Furthermore, the reduced weight of periovarian fat in AE-treated mice was abolished in mice harboring the antibiotic-resistant flora (Fig. S4A), indicating that the reduction in VAT deposits was mediated by depletion of the flora. As previously observed in abx-treated (Savage and Dubos, 1968) and germ-free mice (Schaedler et al., 1965), the weights of the cecum + intestines (small and large bowel) in abx-treated mice were nearly double that of untreated mice (Fig. 3C and D). The increase in cecal and intestinal weight accounted for about 10% of the total body weight and likely contributed to the weight re-gain of abx-treated mice, despite the loss of VAT. The cecal content also showed a darkened color in antibiotic-treated mice compared to untreated mice which was not the result of hematochezia as tests for occult hemoglobin in cecum and large intestine were negative (data not shown). It is known that VAT is preferentially lost when dietary restriction is implemented (Shi et al., 2007), and since the intestinal flora supports the host by breaking down complex dietary fibers that are otherwise not digestible (Bergman, 1990), we hypothesized that depletion of the flora decreased the amount of energy harvested from carbohydrates in the diet. In support of this, AE-treated mice had higher fecal output (Fig. S4B) despite consuming a comparable mass of food (Fig. S4C) compared to mice with an intact flora. To further determine the metabolic phenotype, AE- and untreated mice were singly housed in metabolic cages. The fraction of energy absorbed from the diet was 8% lower in AE-treated mice based on energy intake and energy excreted in feces. (Fig. 3E – G). Antibiotic-treated mice also had a lower energy expenditure compared to untreated mice (Fig 3H, Fig. S4D) which was partly due to less movement (Fig 3I). Reduced nutritional absorption could be a result of disrupted epithelial function in the intestine. However, intestinal epithelial integrity, as measured by leakage of FITC-dextran into the systemic circulation, were not altered in AE-treated mice compared to untreated mice (Fig. S4E). Furthermore, pathology scores for intestinal apoptosis, inflammation, and erosion were not increased in AE-treated mice compared to untreated mice (Fig. S4F). Metabolic profiling also showed that AE-treated transplanted mice had a higher dependence on fat metabolism compared with transplanted mice with an intact flora (Fig. 3J). This was inferred from the respiratory exchange ratio (RER) which varies depending on what fuel is metabolized, with a value of 1 expected when purely carbohydrates are metabolized and a value of 0.7 expected when purely fats are metabolized (Frayn, 1983). The lower RER in AE-treated mice is thus consistent with the notion that antibiotic-treated mice compensated for reduced caloric uptake from carbohydrates by utilizing endogenous fat resulting in reduced VAT.

Fig. 3. Abx-mediated depletion of the intestinal microbiota reduces body fat and caloric uptake from the diet.

Fig. 3

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(A) Weight of untreated (n = 10), AE-treated (n = 8), and VA-treated (n = 10) mice after BMT, relative to day 0. (B) Weight of periovarian fat in untreated (n = 15), AE-treated (n = 14), and VA-treated (n = 10) 28 days after BMT. (C) Representative photographs of intestines and cecum in untreated, AE-treated and VA-treated mice 28 days after BMT. (D) Quantification of weight of intestines + cecum including contents (from duodenum to rectum) of untreated (n = 5), AE-treated (n = 5), and VA-treated (n = 5) mice 28 days after BMT. (E) Energy intake during 24h, (F) energy excreted as feces during 24h, (G) fraction of energy intake absorbed (absorbed energy (ingested calories after subtraction of excreted calories) divided by ingested calories), (H) energy expenditure rate and total energy expenditure during 24h, (I) cumulative and total distance travelled, and (J) respiratory exchange ratio for untreated (n = 10) and AE-treated (n = 10) mice 13 days after BMT. Light and dark cycle indicated by white and grey background, respectively.

Results except (C) and (D) represent at least two independent experiments. * P < 0.05, ** P < 0.01, *** P < 0.001, n.s. – Not significant. Data is presented as mean ± SEM. See also Figure S4.

Sucrose supplementation improves post-transplant lymphopoiesis in mice with an abx-depleted intestinal flora

Dietary restriction, both in experimental models and in human patients with eating disorders such as Anorexia nervosa, has an impact on hematopoiesis, including reduced lymphocyte numbers (Elegido et al., 2017; Tang et al., 2016). To test if reduced uptake of calories in flora-depleted animals contributed to impaired post-BMT hematopoiesis, we supplemented the drinking water with 5% sucrose (Fig. 4A). Although not the form of energy usually provided to the host by the intestinal bacteria, sucrose is a simple carbohydrate absorbed directly by the host without the aid of the flora. This concentration has been previously shown to not induce a preference for the water and to maintain consumption of chow relatively constant (Lewis et al., 2005), with an expected supplementation of approximately 0.9 kcal/mouse/day (about 10% of daily caloric intake). Sucrose supplementation increased peripheral WBC counts after BMT, and particularly increased the low lymphocyte count in AE-treated mice (Fig. 4B). Sucrose supplementation also increased bone marrow cellularity, bone marrow B cell and myeloid frequency although this did not reach statistical significance (Fig. 4C). In addition, sucrose supplemented AE-treated mice had normalized thymocyte counts and showed a trend of increased periovarian VAT mass compared to non-supplemented AE-treated mice (Fig. 4D and E). Sucrose supplementation did not alter fecal bacterial abundance or significantly shift the composition of the remaining intestinal flora in AE-treated mice (Fig. 4F and G, Fig S4G). Taken together, these data suggest that sucrose supplementation can compensate for decreased post-transplant lymphopoiesis due to loss of intestinal flora.

Fig. 4. Sucrose supplementation rescues impaired hematopoietic recovery after BMT in mice with a depleted flora.

Fig. 4

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(A) Experimental outline of sucrose supplementation in water and BMT. (B) WBC, RBC, PLT, LYMPH, NEUT, and MONO counts after BMT, (C) total bone marrow cellularity and composition, (D) total thymocyte count, and (E) weight of periovarian fat 28 days after BMT in untreated and AE-treated mice with and without 5% sucrose in drinking water (n = 10 per group). (F) Quantification of bacterial 16S rRNA and (G) principal components 1 and 2 based on weighted normalized Unifrac analysis of 16S Operational Taxonomic Unit (OTU) abundance in fecal samples 28 days after BMT from untreated and AE-treated mice with and without administration of 5% sucrose in drinking water (untreated mice without sucrose administration, n = 3, all other groups n = 5). Numbers within brackets are percent variation explained by the component.

Shaded areas in (B) indicate normal ranges. Results except (F) and (G) represent two independent experiments. * P < 0.05, ** P < 0.01, *** P < 0.001, n.s. – Not significant. Data is presented as mean ± SEM. See also Figure S4.

The connection between intestinal flora disruption, reduction in body fat, and impaired post-transplant hematopoiesis is dose dependent

Having demonstrated that two different flora-depleting abx regimens impair post-transplant hematopoiesis (Fig. 1 and Fig. 2), at least partly by reducing caloric uptake from the diet (Fig. 3 and Fig. 4), we asked whether perturbations less severe than near-decontamination would also affect hematopoietic recovery. To generate varying degrees of microbiota perturbation, we treated mice with four abx drugs with different spectra of activity; metronidazole, streptomycin, ampicillin, and aztreonam (Fig. 5A). These drugs induced variegated injury to the microbiota as assessed by their effect on total bacterial abundance (16S rRNA qPCR) and diversity (Shannon index) (Fig. 5B and Fig. S5A). Despite its anaerobic-targeted spectrum, metronidazole did not induce major changes to the bacterial diversity or load except for an increase in Faecalibaculum (Fig. S5A). Streptomycin- and aztreonam-treated mice showed a trend of reduced bacterial diversity but had bacterial loads similar to those of untreated mice, while ampicillin reduced the bacterial load dramatically (Fig. 5B). In agreement with a previous study (Berg, 1981), ampicillin had the most severe effects on the intestinal flora and suppressed most intestinal bacterial strains. Therefore, the Shannon diversity index of ampicillin-treated mice was relatively similar to that of untreated mice, since this metric relies on relative bacterial abundance and is insensitive to absolute bacterial load (Fig. 5B). To obtain a parameter that summarized the degree of microbiota injury based on loss of diversity and reduction in total bacterial abundance, each fecal sample was assigned a score derived from both the bacterial diversity and load (Intestinal microbiota score = Shannon Index × 16S rRNA copies per g feces) (Fig. 5C). The degree of intestinal flora integrity 14 days after BMT as assessed by this score correlated positively both with amount of periovarian VAT and post-transplant hematopoietic cell levels including bone marrow cellularity and number of thymocytes 28 days after BMT (Fig. 5D), further supporting the connection between the intestinal flora, nutrition, and post-transplant hematopoiesis. Total bacterial abundance as an individual value also correlated positively to immune reconstitution parameters while bacterial diversity alone did not (Fig. S5B).

Fig. 5. Dose dependent relationship between intestinal flora injury and post-BMT hematopoiesis.

Fig. 5

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(A) Schematic outline of abx treatment and BMT. (B) Fecal bacterial diversity (Shannon index) and fecal bacterial abundance (16S rRNA copies) in samples collected on day 0 and day 14 after BMT from untreated mice (No abx, n = 6), and mice treated with streptomycin (n = 8), metronidazole (n = 10), aztreonam (n = 6), or ampicillin (n = 4). Ellipses show 95% confidence intervals. (C) Intestinal integrity described as an intestinal microbiota score = Shannon index × fecal bacterial abundance for samples shown in (B). (D) Pearson correlation of intestinal microbiota score 14 days after BMT, weight of periovarian fat, bone marrow-, and thymus cellularity in untreated mice (No abx, n = 3) and mice treated with streptomycin (n = 4), metronidazole (n = 5), aztreonam (n = 4), and ampicillin (n = 3).

Results represent one experiment. * P < 0.05, ** P < 0.01. Data is presented as mean ± SEM. See also Figure S5.

Ampicillin and aztreonam-treatment caused comparable reductions in periovarian VAT and post-transplant lymphopoiesis as measured by thymic cellularity (Fig. 5D and Fig. S5C). Interestingly, despite having similar effects on the host animal, the intestinal microbiota phenotype induced by ampicillin was a near-total gut decontamination while aztreonam induced only a modest reduction in total bacterial abundance with a domination of Blautia (Fig. S5A). Interestingly, the genomes of Blautia species encode relatively few enzymes active against the main dietary polysaccharides derived from plant cells (El Kaoutari et al., 2013) (Fig. S5D). In contrast, genera whose genomic repertoires encode many carbohydrate-active enzymes (CAZymes), e.g. Clostridium and Muribaculum, were dramatically depleted after aztreonam treatment (Fig S5E).

Together these results suggest that the disruption of intestinal flora can influence immune reconstitution not only in a binary fashion (severe injury vs. healthy) but rather in a dose-dependent fashion based both on the degree of flora injury and the specific spectrum of the bacterial species affected.

Abx-mediated depletion of the intestinal microbiota under steady-state conditions reduces lymphocyte numbers

Since abx-mediated flora depletion had a strong effect on the hematopoietic phenotype after BMT, we also administered the same AE regimen to non-transplanted mice to determine effects of depletion of the flora on steady-state hematopoiesis. To discriminate between effects of the abx drugs and microbiota-dependent effects we treated both SPF and germ-free (GF) mice (Fig. 6A). AE-treatment for 5 weeks did not alter the total bone marrow cellularity in either SPF or GF mice (Fig 6B, Fig. S6A), but reduced the numbers of bone marrow B cells and thymocytes in SPF mice compared to untreated mice (Fig. 6C and D, Fig. S6B and C). These effects were not seen in the AE-treated GF mice, indicating that it was caused by depletion of the flora. Similar to the transplanted mice, the SPF steady-state AE-treated mice had reduced periovarian VAT after 5 weeks of treatment compared to untreated mice (Fig. 6E), indicating that the phenomenon of abx-induced reduction in body fat was not exclusive to the post-transplant setting. Furthermore, analysis of food intake and fecal output showed comparable food intake and higher fecal output in AE-treated mice compared to untreated controls (Fig. S6D and E), similar to the results obtained for the transplanted mice (Fig. S4B and C).

Fig. 6. Intestinal flora depletion affects steady-state hematopoiesis.

Fig. 6

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(A) Outline of abx-treatment of mice at steady-state without BMT. (B) Bone marrow cellularity, (C) number of bone marrow B cells (B220+), and (D) number of thymocytes in untreated specific pathogen free (SPF) mice (n = 14), AE-treated SPF mice (n = 15), untreated germ-free (GF) mice (n = 10) and AE-treated GF mice (n = 12) after 35 days of antibiotic treatment. (E) Weight of periovarian fat in untreated SPF mice (n = 10) and AE-treated SPF mice (n = 10) after 35 days of antibiotic treatment.

Results represent at least two independent experiments. * P < 0.05, *** P < 0.001, n.s. – Not significant. Data is presented as mean ± SEM. See also Figure S6.

Discussion

In this study, we sought to determine the role of the intestinal flora on hematopoietic recovery after syngeneic BMT using an abx-treatment mouse model. Our results show that depletion of the intestinal flora led to a profound reduction in lymphocyte counts, reduced neutrophil counts, and an impaired capacity to clear systemic infection after BMT. At the same time, initial homing of stem cells, expansion of stem- and progenitor compartments and erythroid differentiation were largely unaffected by the presence of an intact intestinal flora. That intestinal bacteria prime myeloid differentiation and improve clearance of pathogens is in line with several previous reports (Balmer et al., 2014; Clarke et al., 2010; Deshmukh et al., 2014; Khosravi et al., 2014; Tada et al., 1996), and reduced lymphocyte numbers in GF or abx-treated mice have also been described (Josefsdottir et al., 2016; Mazmanian et al., 2005). Our results show no effect or only minor effects on the hematopoietic stem- and progenitor compartments in mice with a depleted intestinal flora, which is in contrast to two studies regarding HSCs in GF and abx-treated mice compared to mice with an intact flora (Iwamura et al., 2017; Josefsdottir et al., 2016). The differences between these studies and ours might be explained by a) the use of a BMT model in our study, b) the use of germ-free mice (Iwamura et al., 2017) vs. abx-treated mice (our study) and c) differences in abx regimens (Josefsdottir et al., 2016). We could also show that depletion of the intestinal flora sensitized mice to a semi-lethal dose of radiation which is in line with previously published work demonstrating that microbiota-derived compounds can protect against irradiation-induced hematopoietic injury (Ainsworth and Mitchell, 1967; Brook and Ledney, 1994; Burdelya et al., 2008; Liu et al., 2015; Smith et al., 1958).

We observed reduced caloric uptake from the diet as a consequence of abx treatment of mice. This is in line with previous observations that dietary energy absorption increases upon bacterial colonization of GF mice (Backhed et al., 2004). A recent study also reported that intestinal bacteria stimulate adsorption of dietary lipids (Wang et al., 2017). Caloric restriction results in lymphopenia both in mice and humans (Elegido et al., 2017; Tang et al., 2016) and we were able to correct impaired post-BMT lymphopoiesis by supplementation of sucrose in the drinking water. Supplementation with this simple sugar rescued peripheral, thymic, and bone marrow lymphocyte counts post-BMT in abx-treated mice. The reduced numbers of neutrophils and bone marrow myeloid cells were only partly rescued by sucrose supplementation, indicating that other mechanisms besides reduced energy extraction induced by flora depletion were involved as well. These might involve the presence of microbial associated molecular patterns (MAMP) which have been reported to be necessary for the maintenance of bone marrow-derived myeloid cells (Khosravi et al., 2014) or gut-microbiota derived peptidoglycans that can modulate neutrophil function (Clarke et al., 2010). The intestinal flora may contribute to energy utilization either by metabolizing complex carbohydrates into simple sugars that the host can directly utilize (Bergman, 1990) and/or by modulating absorptive and functional properties of the intestines such as gut-transit time (Wichmann et al., 2013). Although we observed only a relatively minor reduction in energy extraction efficiency of about 8%, when compounded over the 28-day treatment period it is likely that this contributes to the loss of VAT after abx treatment as shown by others (Suarez-Zamorano et al., 2015). Harvest of nutrients from the diet is a well-recognized feature of the symbiotic relationship between microbiota and host across evolution (Backhed et al., 2005; Wong et al., 2014) and here we extend the relevance of this relationship to systemic immune recovery.

Patients undergoing allogeneic BMT have prolonged and profound nutritional alterations owing to the gastrointestinal and oral mucosal toxicities of pre-transplant conditioning regimens (Baumgartner et al., 2017; Kyle et al., 2005; Lemal et al., 2015; Papadopoulou et al., 1996), and impaired nutritional status after BMT is a negative prognostic marker of overall survival after BMT (Schulte et al., 1998). We conclude that apart from specific and direct effects of the microbiota on immuno-hematopoiesis as previously described (Balmer et al., 2014; Clarke et al., 2010; Khosravi et al., 2014; Mazmanian et al., 2005), our study demonstrates that energy harvest from the diet is a critical mechanism by which the intestinal flora contributes to hematopoiesis after BMT. This observation may help inform the development of strategies to improve immune reconstitution after transplantation.

STAR methods

Contact for reagent and resource sharing

Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Marcel R.M. van den Brink (vandenbm@mskcc.org).

Experimental model and subject details

Mice

Mice were kept under SPF conditions except where germ-free conditions are specified and maintained on a 12 hour light/dark cycle with unlimited access to water and food (5053 Rodent Diet 20, PicoLab). Female mice were used to facilitate randomized mixing between the experimental groups prior to every experiment. C57BL6/J mice were used as host and bone marrow cells for transplantation were prepared from female B6.SJL-PtprcaPepcb/Boy (Ly5.1) or female 129S1/SvImJ mice. All mice were obtained from Jackson Laboratories and used at 8–10 weeks of age. All experiments were performed using protocols approved by the Memorial Sloan-Kettering Cancer Center (MSKCC) Institutional animal care and use committee (IACUC).

Microbial strains

Listeria monocytogenes 10403s was cultured in suspension in BBL brain-heart-infusion media or on BBL brain-heart-infusion agar plates for enumeration. Listeria monocytogenes at log-phase growth was mixed with autoclaved glycerol (20% final concentration) and kept as stocks at −80°C.

Method details

Radiation and bone marrow transplantation

Mice were given a split 1100cGy radiation dose and administered 5 × 106 bone marrow cells via tail vein injection. A single 750cGy radiation dose was used to test endogenous reconstitution. For analysis of homing, 5 × 106 CFSE-stained bone marrow cells or 2 × 104 Lineage−Sca1+ckit+ cells sorted from the bone marrow of GFP+ B6 mice (C57BL/6-Tg(UBC-GFP)30Scha/J) were injected via the tail vein and organs harvested 16h later. Complete blood counts from sub-mandibular bleeds were analyzed using a ProCyte Dx Hematology Analyzer (IDEXX Laboratories). Normal CBC ranges were based on data from the C57BL/6 North American Colonies (Charles-River, 2012).

Drug treatments and sucrose supplementation

Ampicillin 0.5g/L and enrofloxacin 0.25g/L or vancomycin 0.5g/L and amikacin 0.5g/L, were given in the drinking water starting 5 days before BMT and throughout the experiment. Metronidazole 0.5g/L, streptomycin 5g/L, or aztreonam 0.5g/L were started 9 days before BMT and administered throughout the experiment. Ampicillin 0.5g/L and enrofloxacin 0.25g/L treatment without BMT was administered in the drinking water for 35 days. Abx water was changed at least every 5–7 days. Sucrose was supplemented (50g/L) in drinking water and changed every 2–3 days.

Listeria infection

Abx treatment was stopped 3 days prior to infection to allow for washout of the abx. Cultured Listeria monocytogenes 10403s was diluted in PBS and 5 ×103 CFUs were administered intravenously via tail vein. Infected mice were euthanized 3 days later, spleen and liver mashed through a 100 μm cell strainer in PBS+0.05%Triton, and plated in serial dilutions on brain-heart-infusion agar plates to assess bacterial load.

Fecal microbiota transfer (FMT)

Cecal content solution for FMT was prepared from untreated C57BL6/J or from Myd88KO mice under continuous amoxicillin treatment (Caballero et al., 2017). The cecum was placed in an anaerobic chamber and its contents were dissolved in 6ml PBS. Recipient mice were given ampicillin 0.5g/L and enrofloxacin 0.25g/L in the drinking water for 72h and abx were removed 24h before FMT was administered as a single dose of 200μl cecal content solution by oral gavage.

16S sequencing and quantitative PCR (qPCR)

DNA from fecal pellets from individual mice was purified using phenol-chloroform extraction with mechanical disruption (bead beating) based on a previously described protocol (Turnbaugh et al., 2009). Bacterial DNA was analyzed using the Illumina MiSeq platform to sequence the V4-V5 region of the 16S rRNA gene. Sequence data were compiled and processed using mothur and screened and filtered for quality (Schloss et al., 2011). Operational taxonomic units (OTUs) were classified to the genus level using NCBI blastn and the alignment hit with the highest score. Principal component analysis was performed upon a weighted and normalized UniFrac (Lozupone et al., 2006) distance matrix of OTU abundance in R software. Abundance of total 16S copies were analyzed by qPCR using SyberGreen mastermix, 16S_qPCR-F and 16S_qPCR-R oligonucleotides on a StepOnePlus RealTime PCR System (Applied Biosystems) with a standard curve based on serial dilutions of a Blautia 16S rRNA sequence cloned into the pcDNA4 plasmid.

Flow cytometry

Bone marrow cells, thymocytes, splenocytes, or peripheral blood cells were suspended in PBS with 2mM EDTA and 5% FCS. Red cells were lysed from bone marrow and peripheral blood cells using BD Pharm Lyse. Mature blood cells were characterized in peripheral blood or bone marrow using CD4-BV711, CD8-BV785, B220-PE, Gr1-APC, Mac1-PE-Cy7, CD45.1-BV650, and CD45.2 PerCp-Cy5.5. Bone marrow stem cells and progenitor cells were characterized using Sca1-PE-Cy7, ckit-APC, IL7Ra-APC-e780, CD48-FITC, CD150-PerCp-Cy5.5, CD34-Alexa-700, CD16/CD32-eFlour450 and a biotin-conjugated lineage cocktail containing antibodies against Gr1, CD3, NK1.1, CD11b, CD11c, CD8, CD4, Ter119, and CD19, and stained secondary with Streptavidin-Qdot605. Thymocytes were analyzed using CD4-BV711, CD8-BV785, CD25-PE-Cy7, and CD44-eFlour450. B cell subsets in bone marrow were characterized using B220-BV650, CD43-FITC, IgM-Pe-Cy7, IgD-Alexa700, Ly-51-PE and CD24/HSA-Biotin secondary stained with Streptavidin-BV785. B cell subsets in the spleen were analyzed using CD19-APC, CD93-FITC, IgM-Pe-Cy7, and CD21/35-APC-Cy7. DAPI was used to exclude dead cells. Stained cells were analyzed on a BD LSR II and data analyzed using FlowJo software.

Colony forming assay and bone marrow histology

Myeloid colony forming assays were performed by plating 2 × 104 bone marrow cells, counted after lysis of red cells, in duplicates in methyl cellulose media supplemented with growth factors. Colonies were counted 12 days later. Morphologic bone marrow assessment was done on Wright-Giemsa-stained bone marrow brushings using an Olympus BX46 microscope (40× objective, 400× magnification, aperture 0.75). Photos were acquired with an Olympus DP21 camera and Olympus DP21-CB software (v. 02.01.01.93).

FITC-dextran

Mucosal integrity was assessed by measuring FITC-dextran intestinal permeability. For this, mice were fasted for 4h prior to 4kDa FITC-Dextran (0.4mg/g body weight) (Sigma-Aldrich, St. Louis, MO) oral gavage. Blood was collected 4hrs later and plasma FITC levels were determined using an Infinite M1000 PRO fluorescence spectrophotometer (Tecan) at 485 (excitation) and 535 nm (emission) along with a standard curve.

Metabolic analyses

Animals were individually housed in a temperature controlled Promethion Metabolic Screening System (Sable Systems International, NV) where indirect calorimetry was performed to assess rates of oxygen consumption and carbon dioxide production. Food intake was acquired gravimetrically and ambulatory activity was acquired using a laser matrix. Mice were acclimated to this environment on a 12 hr light/dark cycle for 48 hrs before the 24 hr recording period began. The respiratory exchange ratio (RER) was calculated as the ratio of carbon dioxide production to oxygen consumption, and energy expenditure was calculated using the Weir equation (Weir, 1949). Fecal bomb calorimetry was performed on feces collected during the 24 hour metabolic cage recording period, dehydrated in an oven at 60C for 48 hrs, then combusted in technical duplicates with a Parr 6725 Semimicro Calorimeter to determine gross caloric density. When not analyzed in metabolic cages, food intake and fecal output were analyzed in cages holding 4–5 mice each and averaged per mouse. Food intake was analyzed by daily weighing of food and fecal output was assessed by collection in 2h-intervals.

Quantification and statistical analysis

Graphpad Prism software and R were used for graphical presentation and statistical calculations. Two-sided Mann-Whitney U-test was used to compare the means of two groups and two-sided Kruskal-Wallis H-test with Dunn’s multiple comparisons test was used to compare the means of more than two groups. Pearson correlation was used to test correlation between two parameters and Log-rank test was used to compare survival between groups. P < 0.05 was considered statistically significant and data are presented as mean±S.E.M. Number of biological replicates (n) and number of independent experiments are indicated in the figure legends.

Data and software availability

The 16S rRNA sequencing data have been deposited in the European Nucleotide Archive (ENA) with study accession number PRJEB24887.

Supplementary Material

supplement

Highlights.

  • Intestinal microbiota depletion impairs hematopoiesis after bone marrow transplantation

  • Intestinal flora depletion decreases energy harvest and reduces visceral adipose tissue

  • Caloric supplementation rescues impaired hematopoiesis in microbiota-depleted mice

  • The effects of intestinal flora disruption are dose dependent

Acknowledgments

We thank Dr. Julie R. White and coworkers at the MSKCC Center for Comparative Medicine and Pathology for pathology evaluations and Dr. Antonio Gomes for excellent computational work. This work was supported by National Institutes of Health (NIH), National Cancer Institute award number P01-CA023766 (M.R.M.v.d.B.) and Project 4 of P01-CA023766 (M.R.M.v.d.B.), NIH, National Heart, Lung, and Blood Institute award numbers R01-HL069929 (M.R.M.v.d.B.) and R01 HL124112 (R.J.R.), NIH, National Institute of Allergy and Infectious Diseases award number R01–AI100288 (M.R.M.v.d.B.), NIH, National Institute of Diabetes and Digestive and Kidney Diseases award numbers DK048873, DK056626, DK103046 (D.E.C.). Support was also received from The Lymphoma Foundation, The Susan and Peter Solomon Divisional Genomics Program, P30 CA008748 Memorial Sloan Kettering Cancer Center Support Grant/Core Grant, the Parker Institute for Cancer Immunotherapy at Memorial Sloan Kettering Cancer Center, the Cancer Prevention and Research Institute of Texas grant RR160089 (R.J.R.), Seres Therapeutics (J.U.P., R.R.J., and M.R.M.v.d.B.), the Swedish Research Council 2016-00149 (A.S.), the Swedish Society for Medical Research P14-0090 (A.S.), and the Swedish Society of Medicine SLS-499181 (A.S.).

Footnotes

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Author Contributions

Conceptualization, A.S., J.U.P., R.R.J., and M.R.M.v.d.B. Investigation A.S., M.b.d.S., A.E.S., C.S.T., A.L., C.J.B., C.D.H., M.D., J.R.C., A.P., Y.S., B.D., E.V., J.J.T., L.J., H.J., S.L., and O.S. Resources, E.G.P., D.E.C. Writing – Original Draft, A.S., Writing – Review & Editing, J.U.P. R.R.J., and M.R.M.v.d.B. Supervison, R.R.J. and M.R.M.v.d.B.

Declaration of Interests

M.R.M.v.d.B. and R.R.J. are members of the scientific advisory board at Seres Therapeutics.

References

  1. Ainsworth EJ, Mitchell FA. Postirradiation leukocyte patterns in mice and dogs treated with endotoxin. USNRDL-TR-67-23. Res Dev Tech Rep. 1967:1–20. [PubMed] [Google Scholar]
  2. Backhed F, Ding H, Wang T, Hooper LV, Koh GY, Nagy A, Semenkovich CF, Gordon JI. The gut microbiota as an environmental factor that regulates fat storage. Proc Natl Acad Sci U S A. 2004;101:15718–15723. doi: 10.1073/pnas.0407076101. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Backhed F, Ley RE, Sonnenburg JL, Peterson DA, Gordon JI. Host-bacterial mutualism in the human intestine. Science. 2005;307:1915–1920. doi: 10.1126/science.1104816. [DOI] [PubMed] [Google Scholar]
  4. Balmer ML, Schurch CM, Saito Y, Geuking MB, Li H, Cuenca M, Kovtonyuk LV, McCoy KD, Hapfelmeier S, Ochsenbein AF, et al. Microbiota-derived compounds drive steady-state granulopoiesis via MyD88/TICAM signaling. J Immunol. 2014;193:5273–5283. doi: 10.4049/jimmunol.1400762. [DOI] [PubMed] [Google Scholar]
  5. Baumgartner A, Bargetzi A, Zueger N, Bargetzi M, Medinger M, Bounoure L, Gomes F, Stanga Z, Mueller B, Schuetz P. Revisiting nutritional support for allogeneic hematologic stem cell transplantation-a systematic review. Bone Marrow Transplant. 2017;52:506–513. doi: 10.1038/bmt.2016.310. [DOI] [PubMed] [Google Scholar]
  6. Belkaid Y, Hand TW. Role of the microbiota in immunity and inflammation. Cell. 2014;157:121–141. doi: 10.1016/j.cell.2014.03.011. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Berg RD. Promotion of the translocation of enteric bacteria from the gastrointestinal tracts of mice by oral treatment with penicillin, clindamycin, or metronidazole. Infect Immun. 1981;33:854–861. doi: 10.1128/iai.33.3.854-861.1981. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Bergman EN. Energy contributions of volatile fatty acids from the gastrointestinal tract in various species. Physiol Rev. 1990;70:567–590. doi: 10.1152/physrev.1990.70.2.567. [DOI] [PubMed] [Google Scholar]
  9. Brook I, Ledney GD. Effect of antimicrobial therapy on the gastrointestinal bacterial flora, infection and mortality in mice exposed to different doses of irradiation. J Antimicrob Chemother. 1994;33:63–72. doi: 10.1093/jac/33.1.63. [DOI] [PubMed] [Google Scholar]
  10. Burdelya LG, Krivokrysenko VI, Tallant TC, Strom E, Gleiberman AS, Gupta D, Kurnasov OV, Fort FL, Osterman AL, Didonato JA, et al. An agonist of toll-like receptor 5 has radioprotective activity in mouse and primate models. Science. 2008;320:226–230. doi: 10.1126/science.1154986. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Caballero S, Kim S, Carter RA, Leiner IM, Susac B, Miller L, Kim GJ, Ling L, Pamer EG. Cooperating Commensals Restore Colonization Resistance to Vancomycin-Resistant Enterococcus faecium. Cell Host Microbe. 2017;21:592–602 e594. doi: 10.1016/j.chom.2017.04.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Charles-River. C57BL/6 Mouse Hematology North American Colonies. 2012 http://www.criver.com, accessed 2017-07-06.
  13. Chusyd DE, Wang D, Huffman DM, Nagy TR. Relationships between Rodent White Adipose Fat Pads and Human White Adipose Fat Depots. Front Nutr. 2016;3:10. doi: 10.3389/fnut.2016.00010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Clarke TB, Davis KM, Lysenko ES, Zhou AY, Yu Y, Weiser JN. Recognition of peptidoglycan from the microbiota by Nod1 enhances systemic innate immunity. Nat Med. 2010;16:228–231. doi: 10.1038/nm.2087. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Deshmukh HS, Liu Y, Menkiti OR, Mei J, Dai N, O’Leary CE, Oliver PM, Kolls JK, Weiser JN, Worthen GS. The microbiota regulates neutrophil homeostasis and host resistance to Escherichia coli K1 sepsis in neonatal mice. Nat Med. 2014;20:524–530. doi: 10.1038/nm.3542. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. El Kaoutari A, Armougom F, Gordon JI, Raoult D, Henrissat B. The abundance and variety of carbohydrate-active enzymes in the human gut microbiota. Nat Rev Microbiol. 2013;11:497–504. doi: 10.1038/nrmicro3050. [DOI] [PubMed] [Google Scholar]
  17. Elegido A, Graell M, Andres P, Gheorghe A, Marcos A, Nova E. Increased naive CD4+ and B lymphocyte subsets are associated with body mass loss and drive relative lymphocytosis in anorexia nervosa patients. Nutr Res. 2017;39:43–50. doi: 10.1016/j.nutres.2017.02.006. [DOI] [PubMed] [Google Scholar]
  18. Eriksson M, Bolme P. The oral absorption of ampicillin, pivampicillin and amoxycillin in infants and children. Acta Pharmacol Toxicol (Copenh) 1981;49:38–42. doi: 10.1111/j.1600-0773.1981.tb00867.x. [DOI] [PubMed] [Google Scholar]
  19. Frayn KN. Calculation of substrate oxidation rates in vivo from gaseous exchange. J Appl Physiol Respir Environ Exerc Physiol. 1983;55:628–634. doi: 10.1152/jappl.1983.55.2.628. [DOI] [PubMed] [Google Scholar]
  20. Goldberg JD, Zheng J, Ratan R, Small TN, Lai KC, Boulad F, Castro-Malaspina H, Giralt SA, Jakubowski AA, Kernan NA, et al. Early recovery of T-cell function predicts improved survival after T-cell depleted allogeneic transplant. Leuk Lymphoma. 2017:1–13. doi: 10.1080/10428194.2016.1265113. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Guillaume T, Rubinstein DB, Symann M. Immune reconstitution and immunotherapy after autologous hematopoietic stem cell transplantation. Blood. 1998;92:1471–1490. [PubMed] [Google Scholar]
  22. Heinen E. Comparative serum pharmacokinetics of the fluoroquinolones enrofloxacin, difloxacin, marbofloxacin, and orbifloxacin in dogs after single oral administration. J Vet Pharmacol Ther. 2002;25:1–5. doi: 10.1046/j.1365-2885.2002.00381.x. [DOI] [PubMed] [Google Scholar]
  23. Honorato RV. CAZy-parser a way to extract information from the Carbohydrate-Active enZYmes Database. The Journal of Open Source Software. 2016;1 [Google Scholar]
  24. Ishaqi MK, Afzal S, Dupuis A, Doyle J, Gassas A. Early lymphocyte recovery post-allogeneic hematopoietic stem cell transplantation is associated with significant graft-versus-leukemia effect without increase in graft-versus-host disease in pediatric acute lymphoblastic leukemia. Bone Marrow Transplant. 2008;41:245–252. doi: 10.1038/sj.bmt.1705891. [DOI] [PubMed] [Google Scholar]
  25. Iwamura C, Bouladoux N, Belkaid Y, Sher A, Jankovic D. Sensing of the microbiota by NOD1 in mesenchymal stromal cells regulates murine hematopoiesis. Blood. 2017;129:171–176. doi: 10.1182/blood-2016-06-723742. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Jagannath C, Wells A, Mshvildadze M, Olsen M, Sepulveda E, Emanuele M, Hunter RL, Jr, Dasgupta A. Significantly improved oral uptake of amikacin in FVB mice in the presence of CRL-1605 copolymer. Life Sci. 1999;64:1733–1738. doi: 10.1016/s0024-3205(99)00111-3. [DOI] [PubMed] [Google Scholar]
  27. Josefsdottir KS, Baldridge MT, Kadmon CS, King KY. Antibiotics impair murine hematopoiesis by depleting intestinal microbiota. Blood. 2016 doi: 10.1182/blood-2016-03-708594. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Khosravi A, Yanez A, Price JG, Chow A, Merad M, Goodridge HS, Mazmanian SK. Gut microbiota promote hematopoiesis to control bacterial infection. Cell Host Microbe. 2014;15:374–381. doi: 10.1016/j.chom.2014.02.006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Kim DH, Kim JG, Sohn SK, Sung WJ, Suh JS, Lee KS, Lee KB. Clinical impact of early absolute lymphocyte count after allogeneic stem cell transplantation. Br J Haematol. 2004;125:217–224. doi: 10.1111/j.1365-2141.2004.04891.x. [DOI] [PubMed] [Google Scholar]
  30. Kyle UG, Chalandon Y, Miralbell R, Karsegard VL, Hans D, Trombetti A, Rizzoli R, Helg C, Pichard C. Longitudinal follow-up of body composition in hematopoietic stem cell transplant patients. Bone Marrow Transplant. 2005;35:1171–1177. doi: 10.1038/sj.bmt.1704996. [DOI] [PubMed] [Google Scholar]
  31. Lemal R, Cabrespine A, Pereira B, Combal C, Ravinet A, Hermet E, Bay JO, Bouteloup C. Could enteral nutrition improve the outcome of patients with haematological malignancies undergoing allogeneic haematopoietic stem cell transplantation? A study protocol for a randomized controlled trial (the NEPHA study) Trials. 2015;16:136. doi: 10.1186/s13063-015-0663-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Lewis SR, Ahmed S, Dym C, Khaimova E, Kest B, Bodnar RJ. Inbred mouse strain survey of sucrose intake. Physiol Behav. 2005;85:546–556. doi: 10.1016/j.physbeh.2005.06.003. [DOI] [PubMed] [Google Scholar]
  33. Liu W, Chen Q, Wu S, Xia X, Wu A, Cui F, Gu YP, Zhang X, Cao J. Radioprotector WR-2721 and mitigating peptidoglycan synergistically promote mouse survival through the amelioration of intestinal and bone marrow damage. J Radiat Res. 2015;56:278–286. doi: 10.1093/jrr/rru100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Lombard V, Golaconda Ramulu H, Drula E, Coutinho PM, Henrissat B. The carbohydrate-active enzymes database (CAZy) in 2013. Nucleic Acids Res. 2014;42:D490–495. doi: 10.1093/nar/gkt1178. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Lozupone C, Hamady M, Knight R. UniFrac–an online tool for comparing microbial community diversity in a phylogenetic context. BMC Bioinformatics. 2006;7:371. doi: 10.1186/1471-2105-7-371. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Luo Y, Chen GL, Hannemann N, Ipseiz N, Kronke G, Bauerle T, Munos L, Wirtz S, Schett G, Bozec A. Microbiota from Obese Mice Regulate Hematopoietic Stem Cell Differentiation by Altering the Bone Niche. Cell Metab. 2015;22:886–894. doi: 10.1016/j.cmet.2015.08.020. [DOI] [PubMed] [Google Scholar]
  37. Maury S, Mary JY, Rabian C, Schwarzinger M, Toubert A, Scieux C, Carmagnat M, Esperou H, Ribaud P, Devergie A, et al. Prolonged immune deficiency following allogeneic stem cell transplantation: risk factors and complications in adult patients. Br J Haematol. 2001;115:630–641. doi: 10.1046/j.1365-2141.2001.03135.x. [DOI] [PubMed] [Google Scholar]
  38. Mazmanian SK, Liu CH, Tzianabos AO, Kasper DL. An immunomodulatory molecule of symbiotic bacteria directs maturation of the host immune system. Cell. 2005;122:107–118. doi: 10.1016/j.cell.2005.05.007. [DOI] [PubMed] [Google Scholar]
  39. Michelis FV, Messner HA, Loach D, Uhm J, Gupta V, Lipton JH, Seftel MD, Kuruvilla J, Kim DD. Early lymphocyte recovery at 28 d post-transplant is predictive of reduced risk of relapse in patients with acute myeloid leukemia transplanted with peripheral blood stem cell grafts. Eur J Haematol. 2014;93:273–280. doi: 10.1111/ejh.12338. [DOI] [PubMed] [Google Scholar]
  40. Papadopoulou A, Lloyd DR, Williams MD, Darbyshire PJ, Booth IW. Gastrointestinal and nutritional sequelae of bone marrow transplantation. Arch Dis Child. 1996;75:208–213. doi: 10.1136/adc.75.3.208. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Peled JU, Devlin SM, Staffas A, Lumish M, Khanin R, Littmann ER, Ling L, Kosuri S, Maloy M, Slingerland JB, et al. Intestinal Microbiota and Relapse After Hematopoietic-Cell Transplantation. J Clin Oncol. 2017 doi: 10.1200/JCO.2016.70.3348. JCO2016703348. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Porrata LF, Markovic SN. Timely reconstitution of immune competence affects clinical outcome following autologous stem cell transplantation. Clin Exp Med. 2004;4:78–85. doi: 10.1007/s10238-004-0041-4. [DOI] [PubMed] [Google Scholar]
  43. Savage DC, Dubos R. Alterations in the mouse cecum and its flora produced by antibacterial drugs. J Exp Med. 1968;128:97–110. doi: 10.1084/jem.128.1.97. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Schaedler RW, Dubs R, Costello R. Association of Germfree Mice with Bacteria Isolated from Normal Mice. J Exp Med. 1965;122:77–82. doi: 10.1084/jem.122.1.77. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Schloss PD, Gevers D, Westcott SL. Reducing the effects of PCR amplification and sequencing artifacts on 16S rRNA-based studies. PLoS One. 2011;6:e27310. doi: 10.1371/journal.pone.0027310. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Schloss PD, Westcott SL, Ryabin T, Hall JR, Hartmann M, Hollister EB, Lesniewski RA, Oakley BB, Parks DH, Robinson CJ, et al. Introducing mothur: open-source, platform-independent, community-supported software for describing and comparing microbial communities. Appl Environ Microbiol. 2009;75:7537–7541. doi: 10.1128/AEM.01541-09. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Schulte C, Reinhardt W, Beelen D, Mann K, Schaefer U. Low T3-syndrome and nutritional status as prognostic factors in patients undergoing bone marrow transplantation. Bone Marrow Transplant. 1998;22:1171–1178. doi: 10.1038/sj.bmt.1701502. [DOI] [PubMed] [Google Scholar]
  48. Shi H, Strader AD, Woods SC, Seeley RJ. Sexually dimorphic responses to fat loss after caloric restriction or surgical lipectomy. Am J Physiol Endocrinol Metab. 2007;293:E316–326. doi: 10.1152/ajpendo.00710.2006. [DOI] [PubMed] [Google Scholar]
  49. Small TN, Papadopoulos EB, Boulad F, Black P, Castro-Malaspina H, Childs BH, Collins N, Gillio A, George D, Jakubowski A, et al. Comparison of immune reconstitution after unrelated and related T-cell-depleted bone marrow transplantation: effect of patient age and donor leukocyte infusions. Blood. 1999;93:467–480. [PubMed] [Google Scholar]
  50. Smith WW, Alderman IM, Gillespie RE. Hematopoietic recovery induced by bacterial endotoxin in irradiated mice. Am J Physiol. 1958;192:549–556. doi: 10.1152/ajplegacy.1958.192.3.549. [DOI] [PubMed] [Google Scholar]
  51. Suarez-Zamorano N, Fabbiano S, Chevalier C, Stojanovic O, Colin DJ, Stevanovic A, Veyrat-Durebex C, Tarallo V, Rigo D, Germain S, et al. Microbiota depletion promotes browning of white adipose tissue and reduces obesity. Nat Med. 2015;21:1497–1501. doi: 10.1038/nm.3994. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Tada T, Yamamura S, Kuwano Y, Abo T. Level of myelopoiesis in the bone marrow is influenced by intestinal flora. Cell Immunol. 1996;173:155–161. doi: 10.1006/cimm.1996.0261. [DOI] [PubMed] [Google Scholar]
  53. Tang D, Tao S, Chen Z, Koliesnik IO, Calmes PG, Hoerr V, Han B, Gebert N, Zornig M, Loffler B, et al. Dietary restriction improves repopulation but impairs lymphoid differentiation capacity of hematopoietic stem cells in early aging. J Exp Med. 2016;213:535–553. doi: 10.1084/jem.20151100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Taur Y, Jenq RR, Perales MA, Littmann ER, Morjaria S, Ling L, No D, Gobourne A, Viale A, Dahi PB, et al. The effects of intestinal tract bacterial diversity on mortality following allogeneic hematopoietic stem cell transplantation. Blood. 2014;124:1174–1182. doi: 10.1182/blood-2014-02-554725. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Tedesco F, Markham R, Gurwith M, Christie D, Bartlett JG. Oral vancomycin for antibiotic-associated pseudomembranous colitis. Lancet. 1978;2:226–228. doi: 10.1016/s0140-6736(78)91741-5. [DOI] [PubMed] [Google Scholar]
  56. Turnbaugh PJ, Hamady M, Yatsunenko T, Cantarel BL, Duncan A, Ley RE, Sogin ML, Jones WJ, Roe BA, Affourtit JP, et al. A core gut microbiome in obese and lean twins. Nature. 2009;457:480–484. doi: 10.1038/nature07540. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Velders GA, van Os R, Hagoort H, Verzaal P, Guiot HF, Lindley IJ, Willemze R, Opdenakker G, Fibbe WE. Reduced stem cell mobilization in mice receiving antibiotic modulation of the intestinal flora: involvement of endotoxins as cofactors in mobilization. Blood. 2004;103:340–346. doi: 10.1182/blood-2002-07-2270. [DOI] [PubMed] [Google Scholar]
  58. Wang Y, Kuang Z, Yu X, Ruhn KA, Kubo M, Hooper LV. The intestinal microbiota regulates body composition through NFIL3 and the circadian clock. Science. 2017;357:912–916. doi: 10.1126/science.aan0677. [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Weir JB. New methods for calculating metabolic rate with special reference to protein metabolism. J Physiol. 1949;109:1–9. doi: 10.1113/jphysiol.1949.sp004363. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Wichmann A, Allahyar A, Greiner TU, Plovier H, Lunden GO, Larsson T, Drucker DJ, Delzenne NM, Cani PD, Backhed F. Microbial modulation of energy availability in the colon regulates intestinal transit. Cell Host Microbe. 2013;14:582–590. doi: 10.1016/j.chom.2013.09.012. [DOI] [PubMed] [Google Scholar]
  61. Williams JP, Brown SL, Georges GE, Hauer-Jensen M, Hill RP, Huser AK, Kirsch DG, Macvittie TJ, Mason KA, Medhora MM, et al. Animal models for medical countermeasures to radiation exposure. Radiat Res. 2010;173:557–578. doi: 10.1667/RR1880.1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  62. Wong AC, Dobson AJ, Douglas AE. Gut microbiota dictates the metabolic response of Drosophila to diet. J Exp Biol. 2014;217:1894–1901. doi: 10.1242/jeb.101725. [DOI] [PMC free article] [PubMed] [Google Scholar]

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