- Record: found
- Abstract: found
- Article: found
Effects of Parental Omega-3 Fatty Acid Intake on Offspring Microbiome and Immunity
research-article
Publication date (Electronic):
29 January 2014
Journal:
PLoS ONE
Publisher:
Public Library of Science
Read this article at
There is no author summary for this article yet. Authors can add summaries to their articles on ScienceOpen to make them more accessible to a non-specialist audience.
Abstract
The “Western diet” is characterized by increased intake of saturated and omega-6 (n−6) fatty acids with a relative reduction in omega-3 (n−3) consumption. These fatty acids can directly and indirectly modulate the gut microbiome, resulting in altered host immunity. Omega-3 fatty acids can also directly modulate immunity through alterations in the phospholipid membranes of immune cells, inhibition of n−6 induced inflammation, down-regulation of inflammatory transcription factors, and by serving as pre-cursors to anti-inflammatory lipid mediators such as resolvins and protectins. We have previously shown that consumption by breeder mice of diets high in saturated and n−6 fatty acids have inflammatory and immune-modulating effects on offspring that are at least partially driven by vertical transmission of altered gut microbiota. To determine if parental diets high in n−3 fatty acids could also affect offspring microbiome and immunity, we fed breeding mice an n−3-rich diet with 40% calories from fat and measured immune outcomes in their offspring. We found offspring from mice fed diets high in n−3 had altered gut microbiomes and modestly enhanced anti-inflammatory IL-10 from both colonic and splenic tissue. Omega-3 pups were protected during peanut oral allergy challenge with small but measurable alterations in peanut-related serologies. However, n−3 pups displayed a tendency toward worsened responses during E. coli sepsis and had significantly worse outcomes during Staphylococcus aureus skin infection. Our results indicate excess parental n−3 fatty acid intake alters microbiome and immune response in offspring.
Related collections
Most cited references22
- Record: found
- Abstract: found
- Article: not found
Probiotics and the gut microbiota in intestinal health and disease.
Melanie Gareau, Philip Sherman, W Allan Walker (2010)
The use of probiotics is increasing in popularity for both the prevention and treatment of a variety of diseases. While a growing number of well-conducted, prospective, randomized, controlled, clinical trials are emerging and investigations of underlying mechanisms of action are being undertaken, questions remain with respect to the specific immune and physiological effects of probiotics in health and disease. This Review considers recent advances in clinical trials of probiotics for intestinal disorders in both adult and pediatric populations. An overview of recent in vitro and in vivo research related to potential mechanisms of action of various probiotic formulations is also considered.
- Record: found
- Abstract: found
- Article: found
Mom Knows Best: The Universality of Maternal Microbial Transmission
Summary The sterile womb paradigm is an enduring premise in biology that human infants are born sterile. Recent studies suggest that infants incorporate an initial microbiome before birth and receive copious supplementation of maternal microbes through birth and breastfeeding. Moreover, evidence for microbial maternal transmission is increasingly widespread across animals. This collective knowledge compels a paradigm shift—one in which maternal transmission of microbes advances from a taxonomically specialized phenomenon to a universal one in animals. It also engenders fresh views on the assembly of the microbiome, its role in animal evolution, and applications to human health and disease. Introduction While the human microbiota comprises only 1–3% of an individual's total body mass, this small percentage represents over 100 trillion microbial cells, outnumbering human cells 10 to 1 and adding over 8 million genes to our set of 22,000 [1],[2]. This complexity establishes a network of interactions between the host genome and microbiome spanning gut development [3], digestion [4],[5], immune cell development [6]–[8], dental health [9],[10], and resistance to pathogens [11],[12]. Recent studies have also provided a greater understanding of how the composition of an individual's microbiota changes throughout development, especially during the first year of life [3],[13]. While the general dogma is that the placental barrier keeps infants sterile throughout pregnancy, increasing evidence suggests that an infant's initial inoculum can be provided by its mother before birth [14]–[18] and is supplemented by maternal microbes through the birthing [19] and breastfeeding [20],[21] processes. While maternal transmission of microbes in humans has attracted considerable attention in the last few years, nearly a century's worth of research is available for vertical transmission of symbionts in invertebrates [22]. Similar to gut bacteria in humans that assist nutrient intake, many insect-associated bacteria function as nutritional symbionts that supplement the nutrient-poor diet of their host with essential vitamins or amino acids [23],[24]. Since these indispensable symbionts cannot live outside of host cells, they cannot be acquired from the environment and are faithfully transferred from mother to offspring [22],[25]. Maternal transmission in invertebrates has been reviewed elsewhere [22],[26],[27], and Box 1 and Box 2 highlight examples of heritable symbioses across invertebrate phyla. Box 1. Examples of Maternal Transmission in Marine Invertebrates Marine Sponges (Phylum Porifera) Sponges are ancient metazoans that evolved over 600 million years ago as one of the first multicellular animals [83]. In marine sponges, a remarkably large consortium of extracellular microbial symbionts thrives within the sponge's mesohyl, a gelatinous connective tissue located between the external and internal cell layers. Many of these bacterial residents are found in diverse species of sponges with nonoverlapping distributions but not in the surrounding seawater [84]–[86]. These “sponge-specific” microbes are hypothesized to have originated from ancient colonization events before the diversification of marine sponges and are maintained as symbionts through vertical transmission [87]. Independent studies have estimated that up to 33 phylogenetically distinct microbial clusters spanning ten bacterial phyla and one archaeal phylum are vertically transmitted in sponges [41],[84],[86],[88]. Both transmission electron microscopy (TEM) and fluorescent in situ hybridization (FISH) studies have confirmed the presence of microorganisms of different shapes and sizes in the oocytes of oviparous sponges [41] and in the embryos of viviparous sponges [43]–[45]. Vesicomyid Clams (Phylum Mollusca) Deep-sea hydrothermal vent communities rely upon chemosynthetic bacteria to harness chemical energy stored in reduced sulfur compounds extruding from the vents. Metazoans that live in this extreme environment harbor chemosynthetic endosymbionts in their tissues that provide most, if not all, of the host's nutrition [89]. Somewhat surprisingly, most invertebrates that live near hydrothermal events acquire their endosymbionts anew from the environment each generation [90],[91], even though chemosynthetic bacteria are crucial for survival in such a harsh habitat. A major exception to this trend is found in the Vesicomyidae family of clams [92]. Vesicomyid clams retain a rudimentary gut and rely primarily on sulfur-oxidizing bacteria sequestered intracellularly within specialized host cells called bacteriocytes in the clam's large, fleshy gills [93]. Vertical transmission via transovarial transmission appears to be the dominant mechanism for maintenance of these thioautotrophic bacterial symbionts given that follicle cells surrounding an oocyte and the oocyte itself are heavily infected with the chemosynthetic bacteria [46],[94]. Box 2. Examples of Maternal Transmission in Terrestrial Invertebrates Insects (Phylum Arthropoda) Insects that thrive on unbalanced diets such as plant sap, blood, or wood depend upon microbial symbionts for the provision of essential amino acids or vitamins lacking in their food source. In turn, hosts provide a wide range of metabolites to their symbionts as well as protection from environmental stressors. This codependence requires faithful transfer of symbionts to all offspring, usually through transovarial transmission [23],[24]. Reproductive parasites, such as the obligate, intracellular bacteria Wolbachia, are also widespread in insects and hijack maternal transmission routes to ensure their spread within an insect population (reviewed in [95],[96]). Pea Aphid (Acrythosiphon pisum) The pea aphid Acrythosiphon pisum (Figure 2A) and its nutritional endosymbiont Buchnera aphidicola are a preeminent example of obligate mutualism in insects. The ancestral Buchnera gammaproteobacteria was acquired by aphids between 160 and 280 million years ago [97] and has since diverged in parallel with its aphid hosts through strict vertical transmission [26],[97]. Buchnera are housed within the cytoplasm of bacteriocytes arranged into dual bacteriome structures located in the aphid body cavity adjacent to the ovaries [98], allowing efficient transfer of Buchnera symbionts to developing oocytes or embryos during the sexual and asexual phases of aphid reproduction, respectively. At the cellular level, symbiont transfer occurs when maternal bacteriocytes release Buchnera symbionts through exocytosis into the extracellular space between the bacteriocyte and oocyte or embryo, which then actively endocytoses the extracellular Buchnera symbionts [51]. Cockroaches (Order Blattodea) Just as insects are morphologically diverse, the mechanisms by which insects transport symbionts to oocytes are highly varied. In cockroaches, Blattabacterium-filled bacteriocyte cells migrate from the abdominal fat body to the distantly located ovarioles where they adhere to the oocyte membrane [99],[100]. Interestingly, the bacteriocytes remain associated with the oocyte for eight to nine days before finally expelling their symbionts through exocytosis. The Blattabacterium cells then squeeze between the follicle cells surrounding the oocyte and are engulfed into the oocyte cytoplasm via endocytosis just prior to ovulation [100]. Whiteflies (Family Aleyrodidae) The whitefly circumvents exocytosis of its intracellular nutritional symbiont, Portiera aleyrodidarum, by depositing entire bacteriocytes into its eggs. These maternal bacteriocytes remain intact yet separate from the developing embryo until the embryonic bacteriomes form, at which point the maternal bacteriocytes deteriorate [22]. Tsetse Flies, Bat Flies, and Louse Flies (Superfamily Hippoboscoidea) Members of the Hippoboscoidea superfamily (Order Diptera) are obligate blood feeders that have developed a unique reproductive strategy termed adenotrophic viviparity that offers a different solution to internal maternal transfer of symbionts. Females of this superfamily develop a single fertilized embryo at a time within their uterus (modified vaginal canal) until it is deposited as a mature third instar larva immediately preceding pupation. During their internal development, the larvae are nourished with milk produced by modified accessory glands that empty into the uterus [101]. The milk primarily consists of protein and lipids [102], but it also serves as a reservoir for maternally transmitted microbial symbionts [103]. For example, the obligate mutualistic symbiont of tsetse flies, Wigglesworthia glossinidia, is absent from the female germ line and surrounding reproductive tissues but is found extracellularly in the female milk glands and is first detected in tsetse offspring once milk consumption begins during the first larval stage [103]. Stinkbugs (Superfamily Pentatomoidea) One of the most common mechanisms of external maternal transmission in insects is that of “egg smearing,” which occurs when a female contaminates the surface of her eggs with symbiont-laden feces during oviposition. Upon hatching, offspring probe or consume the discarded egg shells to acquire the maternal bacteria. This mode of transmission is commonly found in plant-sucking stinkbugs, including the Pentatomidae and Acanthosomatidae families [104]. In the Cynidae family of stinkbugs, along with the Coreidae family of leaf-footed bugs, gut symbionts are transferred maternally via coprophagy, in which offspring consume maternal feces, sometimes directly from the mother's anus [22],[104]. Stinkbugs of the Plataspidae family, on the other hand, have developed a unique mode of transmission via a maternally provided “symbiont capsule” deposited on the underside of the egg mass [56]. These capsules are comprised of bacterial cells dispersed throughout a resin-like matrix surrounded by a brown, cuticle-like envelope that protects the symbionts from environmental stressors such as UV irradiation or dissection [57]. After hatching, plataspid nymphs immediately probe the capsules to ingest the symbionts [56],[59]. European Beewolf (Philanthus triangulum) While nutritional symbionts appear to be the most common type of bacteria transmitted via external maternal transmission in insects, the European beewolf (Philanthus triangulum) instead cultivates a symbiotic bacteria that protects offspring against microbial infection during development. Beewolves are solitary digger wasps that deposit their offspring in moist, underground nests, making them susceptible to fungal and bacterial infections [105]. To combat these pathogens, female beewolves cultivate Streptomyces philanthi bacteria in specialized glands in their antennae, which they copiously spread on the ceiling of the brood cell before oviposition [106]–[108]. After hatching, the larvae take up the bacterial cells and incorporate them into their cocoon that they build before pupation. When adult beewolves emerge from their cocoon in the summer, female beewolves acquire the maternally provided Streptomyces symbiont and house them in the female-specific gland reservoirs along each antenna [108],[109]. By integrating previous studies in invertebrates with recent evidence for maternal microbial transmission in humans and other vertebrates, we contend that maternal provisioning of microbes is a universal phenomenon in the animal kingdom. As a result, a considerable new phase of study in heritable symbiont transmission is underway. Thus, this essay presents current evidence for maternal microbial transmission and provides new insights into its impact on microbiome assembly and evolution, with applications to human health and disease. Internal Maternal Transmission At the turn of the twentieth century, French pediatrician Henry Tissier asserted that human infants develop within a sterile environment and acquire their initial bacterial inoculum while traveling through the maternal birth canal [28]. More than a century later, the sterile womb hypothesis remains dogma, as any bacterial presence in the uterus is assumed to be dangerous for the infant. Indeed, studies of preterm deliveries have found a strong correlation between intrauterine infections and preterm labor, especially when birth occurs less than 30 weeks into the pregnancy [29],[30]. Since preterm birth is the leading cause of infant mortality worldwide [31], much attention has focused on identifying the bacterial culprits responsible for spontaneous preterm labor. Surprisingly, most of the bacteria detected in intrauterine infections are commonly found in the female vaginal tract [29], and risk of preterm birth is markedly increased in women diagnosed with bacterial vaginosis during pregnancy [32]. Interestingly, the vaginal microbial community varies significantly among American women of different ethnicities (Caucasian, African-American, Asian, or Hispanic), with African-American and Hispanic women more likely to have a microbiota traditionally associated with bacterial vaginosis (predominance of anaerobic bacteria over Lactobacillus species) [33] and a higher rate of spontaneous preterm deliveries (reviewed in [34]). While intrauterine infection and inflammation is important in understanding the etiology of preterm birth, relatively few studies have examined the uterine microbiome of healthy, term pregnancies owing to the sterile womb paradigm. Investigations into the potential for bacterial transmission through the placental barrier have detected bacteria in umbilical cord blood [17], amniotic fluid [14],[35], and fetal membranes [35],[36] from babies without any indication of inflammation (Figure 1). Furthermore, an infant's first postpartum bowel movement of ingested amniotic fluid (meconium) is not sterile as previously assumed, but instead harbors a complex community of microbes, albeit less diverse than that of adults [18],[37]. Interestingly, many of the bacterial genera found in the meconium, including Enterococcus and Escherichia, are common inhabitants of the gastrointestinal tract [18],[37]. To test whether maternal gut bacteria can be provisioned to fetuses in utero, Jiménez et al. [18] fed pregnant mice milk inoculated with genetically-labeled Enterococcus faecium and then examined the meconium microbes of term offspring after sterile C-section. Remarkably, E. faecium with the genetic label was cultured from the meconium of pups from inoculated mothers, but not from pups of control mice fed noninoculated milk. Meconium from the treatment group also had a higher abundance of bacteria than that of the control group. Importantly, the study controlled for potential bacterial contamination from contact between skin and the meconium by sampling an internal portion of the meconium [18]. Thus, this study provides foundational evidence for maternal microbial transmission in mammals. 10.1371/journal.pbio.1001631.g001 Figure 1 Sources of microbial transmission in humans from mother to child. Cut-away diagram highlighting the various internal and external sources of maternal microbial transmission as well as the species that are commonly associated with transfer from those regions. Regions discussed include the oral cavity [14],[16], the mammary glands [40],[78],[79], the sebaceous skin surrounding the breast [78],[80], the vaginal tract [19],[33],[73], and the intrauterine environment [14],[15],[17],[18],[29],[35]–[37],[40]. Illustration by Robert M. Brucker. Other than ascension of vaginal microbes associated with preterm births, the mechanisms by which gut bacteria gain access to the uterine environment are not well understood. One possibility is that bacteria travel to the placenta via the bloodstream after translocation of the gut epithelium. While the intestinal epithelial barrier generally prevents microbial entry into the circulatory system, dendritic cells can actively penetrate the gut epithelium, take up bacteria from the intestinal lumen, and transport the live bacteria throughout the body as they migrate to lymphoid organs [38],[39]. Interestingly, microbial translocation may even increase during pregnancy, as one study showed that pregnant mice were 60% more likely to harbor bacteria in their mesenteric lymph node (presumably brought there by dendritic cells) than nonpregnant mice [40]. Bacterial species normally found in the human oral cavity have also been isolated from amniotic fluid and likely enter the bloodstream during periodontal infections, facilitated by gingiva inflammation [14],[16] (Figure 1). Overall, the study of internal maternal transmission of microbes in mammals is in its infancy due to the enduring influence of the sterile womb paradigm and to the ethical and technical difficulties of collecting samples from healthy pregnancies before birth. Thus, we still know very little about the number and identity of innocuous microbes that traverse the placenta, whether they persist in the infant, or whether their presence has long-term health consequences for the child. Similarly, we know almost nothing about nonpathogenic viruses or archaea that may be transferred from mother to child alongside their bacterial counterparts. Fortunately, the advent of culture-independent, high-throughput sequencing will serve as a tremendous resource for this field and will hopefully lead to a characterization of the “fetal microbiome” in utero. Maternal provisioning of microbes to developing offspring is widespread in animals, with evidence of internal microbial transmission in animal phyla as diverse as Porifera [41]–[45] (Box 1), Mollusca [46]–[49] (Box 1), Arthropoda [50]–[52] (Box 2, Figure 2), and Chordata [19],[53],[54] (Box 3, Figure 2). The presence of maternal transmission at the base of the Animalia kingdom and the surprising plasticity by which microbes gain access to germ cells or embryos in these systems signifies that maternal symbiont transmission is an ancient and evolutionarily advantageous mechanism inherent in animals, including humans. Therefore, we can no longer ignore the fact that exposure to microbes in the womb is likely and may even be a universal part of human pregnancy, serving as the first inoculation of beneficial microbes before birth. 10.1371/journal.pbio.1001631.g002 Figure 2 Examples of animals that exhibit microbial maternal transmission. (A) Pea aphid (Acyrthosiphon pisum), photo credit: Whitney Cranshaw, Colorado State University/©Bugwood.org/CC-BY-3.0-US; (B) Domesticated chicken hen (Gallus gallus domesticus), photo credit: Ben Scicluna; (C) Sockeye salmon (Oncorhynchus nerka), photo credit: Cacophony; (D) South American river turtle (Podocnemis expansa), photo credit: Wilfredor. All photos were obtained from Wikimedia Commons (www.commons.wikimedia.org). Box 3. Examples of Maternal Transmission in Vertebrates Aside from studies in human and mouse models, very little is known about maternal transmission of microbial communities in vertebrates, especially outside Class Mammalia. Furthermore, research on vertical transmission in nonmammalians has largely focused on maternally transmitted pathogens, especially in animals of agricultural importance like chickens and fish. Domesticated Chickens (Gallus gallus domesticus) Zoonotic Salmonella infections acquired from contaminated chicken eggs is estimated to cause more than 100,000 illnesses each year in the United States [110]. In addition to horizontal transmission of Salmonella on eggs through surface contamination, direct transovarial transmission also occurs when Salmonella colonizes the reproductive tissues of hens (Figure 2B). Depending on the infection location within the female reproductive tract, the bacteria are deposited into the yolk, albumen, eggshell membrane, and/or eggshell of the developing egg before oviposition (reviewed in [111]). Other poultry pathogens, such as Mycoplasma synoviae in chickens [112] and M. gallisepticum, M. cloacale, and M. anatis in ducks [113], have also been cultured from the yolk of embryonated eggs, though whether commensal flora are incorporated into the egg is not known. Ray-Finned Fish (Class Actinopterygii) Several bacterial pathogens of economically important fish are transmitted transovarially in the egg yolk including Renibacterium salmoninarum, the agent of bacterial kidney disease in salmonids (Figure 2C), and Flavobacterium psychrophilum, which causes bacterial cold water disease in salmonids and rainbow trout fry disease in trout (reviewed in [114]). F. psychrophilum has also been found in ovarian fluid and on the surface of eggs of steelhead trout [115]. Additionally, an obligate, intracellular eukaryotic parasite, Pseudoloma neurophilia, is a common pathogen found in zebrafish (Danio rerio) facilities and has been observed in spores of the ovarian stroma and within developing follicle cells of spawning females, suggesting that it can be vertically transmitted, though it is primarily spread from fish to fish in contaminated water (reviewed in [116]). Turtles (Order Chelonii) The formation of egg components in the uterine tube and uterus of turtles takes approximately two weeks, providing ample opportunity for maternal transmission of intestinal or reproductive microbes to the egg [117]. One study of unhatched (dead) eggs from loggerhead sea turtle (Caretta caretta) nests found several potential pathogens, including Pseudomonas aeruginosa and Serratia marcesans, in fluid from the interior of the eggs, though environmental contamination of the eggs cannot be ruled out [118]. A similar study of eggs from two species of South American river turtles, Podocnemis expansa (Figure 2D) and P. unifilis, identified several Enterobacteriaceae species, including Escherichia coli, Shigella flexneri, and Salmonella cholerasuis, in the eggs but not in the environmental samples taken from the turtle nests [119], suggesting that they may have a maternal origin. In support of this hypothesis, a separate study in green turtles (Chelonia mydas) that collected eggs directly from the maternal cloacal opening during egg laying isolated Pseudomonas, Salmonella, Enterobacter, and Citrobacter from the eggshell, albumen, and yolk. In fact, the yolk was the egg component most heavily infected with bacteria [120]. Altogether, many potentially pathogenic species have been isolated from turtle eggs, but whether these bacteria actually cause disease in turtles or are part of their natural flora remains to be determined. External Maternal Transmission External maternal transmission encompasses any transfer of maternal symbionts to offspring during or after birth. In invertebrates, it is often accomplished by “egg smearing,” in which females coat eggs with microbes as they are deposited [55], or through the provision of a microbe-rich maternal fecal pellet that is consumed by larval offspring upon hatching [56]–[59] (see Box 2). Similarly, human infants are “smeared” with maternal vaginal and fecal microbes as they exit the birth canal [60]–[62] (Figure 1). Several studies have shown that the human neonatal microbiota across all body habitats (skin, oral, nasopharyngeal, and gut) is influenced by their mode of delivery [19],[63]–[65], with infants born vaginally acquiring microbes common in the female vagina while C-section infants display a microbiota more similar to that of human skin [19]. Furthermore, while the microbiota of a vaginally delivered infant clusters with the vaginal bacteria of its mother, the microbiota of C-section babies is no more related to the skin flora of its mother than that of a stranger, indicating that most microbes are transmitted to the neonate from those handling the infant [19]. Importantly, epidemiological data suggest that a Cesarean delivery can have long-term consequences on the health of a child, especially concerning immune-mediated diseases. For example, children born via C-section are significantly more likely to develop allergic rhinitis [66], asthma [66], celiac disease [67], type 1 diabetes [68], and inflammatory bowel disease [69]. These statistics are alarming given that 32.8% of all births in the United States in 2010 were delivered via C-section with similar rates on the rise in most developed countries [70]. The higher rate of immune-mediated diseases in C-section children may indicate that maternally transferred vaginal or fecal microbes are unique in their ability to elicit immune maturation in the neonate. Development of the intestinal mucosa and secondary lymphoid tissues in the gut is contingent upon recognition of microbial components by pattern-recognition receptors on intestinal epithelial cells (reviewed in [71],[72]). It is possible that these receptors cannot properly interact with the community of microbes acquired during Cesarean deliveries, leading to disrupted immune development and an increased risk for immune-mediated disorders in C-section children. Conversely, transmission for thousands of years of vaginal and fecal microbes at birth has likely produced specific human-microbe interactions important for neonatal gut development. In fact, a recent study found that the vaginal microbial community changes during pregnancy, becoming less diverse as the pregnancy progresses [73]; yet, in spite of the general decrease in richness, certain Lactobacillus bacterial species are enriched in the vaginal community during pregnancy and are hypothesized to be important for establishing the neonatal upper GI microbiota after vaginal delivery [73]. Breastfeeding provides a secondary route of maternal microbial transmission as shown in humans (reviewed in [74], Figure 1) and nonhuman primates such as rhesus monkeys [75]. In humans, maternal milk microbes are implicated in infant immune system development [76], resistance against infection [77], and protection against the development of allergies and asthma later in childhood [74]. High-throughput sequencing of breast milk from 16 healthy women identified 100–600 species of bacteria in each sample with nine genera present in every sample: Staphylococcus, Streptococcus, Serratia, Pseudomonas, Corynebacterium, Ralstonia, Propionibacterium, Sphingomonas, and Bradyrhizobiaceae [78]. This “core” milk microbiome represented approximately 50% of all bacteria in each sample, with the other half representing individual variation in microbial composition [78]. A similar study found that the bacterial composition in breast milk changes over time: milk produced immediately after labor harbored more lactic acid bacteria along with Staphylococcus, Streptococcus, and Lactococcus, while breast milk after six months of lactation had a significant increase in typical inhabitants of the oral cavity, such as Veillonella, Leptotrichia, and Prevotella [79], perhaps to prime the infant for the switch to solid food. However, as with any DNA-based, culture-independent study that does not discriminate between live and dead bacteria, the number and identity of bacteria detected in these studies should be interpreted with some caution. Given that milk is only produced temporarily in a woman's life, the origin of milk microbes is still somewhat of a mystery. Breast milk was traditionally thought to be sterile; however, colostrum (the first milk produced after delivery) collected aseptically already harbors hundreds of bacterial species [79]. Breast milk does share many taxa with the microbiota found on sebaceous skin tissue around the nipple [78],[80], and high levels of Streptococcus in breast milk may be a result of retrograde flow from an infant's oral cavity back to the milk ducts during suckling [81] since Streptococcus is the dominant phylotype in infant saliva [82]. However, the presence of anaerobic gut bacteria in human milk suggests that an entero-mammary route of transfer also exists that may utilize phagocytic dendritic cells to traffic gut microbes to the mammary glands, similar to microbial transfer to amniotic fluid as discussed earlier. To support this hypothesis, Perez et al. [40] found identical strains of bacteria in milk cells, blood cells, and fecal samples from lactating women, but more work is needed to directly connect bacterial translocation in the gut to incorporation in breast milk. Overall, maternal transmission of beneficial microbes in humans has widespread relevance for human health. Evolution with these microbes has resulted in our dependence on them for the proper maturation and development of the immune system and gastrointestinal tract. Somewhat paradoxically, modern medicine designed to prevent infant mortality (such as emergency Cesarean sections and formula feeding) has likely contributed to the rise in immune-mediated diseases in developed countries due to the inherent lack of exposure to maternal microbes associated with these practices. Fortunately, biomedicine is also making strides in finding effective probiotic supplements to promote immune development and ameliorate some of the risks that C-section or formula-fed infants face as children and adults. Hopefully, as we gain understanding of the diversity and function of maternally transmitted microbes in humans, more complete and effective probiotic blends will recapitulate the microbial communities found in vaginally delivered, breast-fed infants and restore the microbe-host interactions that humans depend upon for proper development. Conclusions Since the early twentieth century, the study of maternal microbial transmission has focused heavily on animal systems in which maternal transmission maintains sophisticated partnerships with one or two microbial species. However, with the development of high-throughput sequencing technologies, it is now possible to identify entire microbiomes that are transferred from mother to offspring in systems not traditionally considered to exhibit maternal transmission, such as humans. By expanding the definition of maternal transmission to include all internal and external microbial transfers from mother to offspring, we contend that maternal transmission is universal in the animal kingdom and is used to provision offspring with important microbes at birth, rather than leave their acquisition to chance. Finally, with microbes contributing 99% of all unique genetic information present in the human body, maternal microbial transmission should be viewed as an additional and important mechanism of genetic and functional change in human evolution. Similar to deleterious mutations in our genetic code, disruption of maternal microbial acquisition during infancy could “mutate” the composition of the microbial community, leading to improper and detrimental host-microbe interactions during development. Maternal transmission is also a key factor in shaping the structure of the microbiome in animal species over evolutionary time, since microbes that promote host fitness, especially in females, will simultaneously increase their odds of being transferred to the next generation. Thus, whether internal or external, the universality and implications of maternal microbial transmission are nothing short of a paradigm shift for the basic and biomedical life sciences.
- Record: found
- Abstract: found
- Article: found
Regulation of intestinal inflammation by microbiota following allogeneic bone marrow transplantation
Robert Jenq, Carles Ubeda, Ying Taur … (2012)
Regulators of the intestinal flora include diet (Kau et al., 2011), antibiotics (Willing et al., 2011), and, importantly, intestinal inflammation (Sekirov et al., 2010). As a result, the cause–effect relationships between intestinal inflammation and changes in microbiota have been difficult to define (Maloy and Powrie, 2011). The success of allogenic BM transplantation (BMT), a standard therapy for conditions such as hematopoietic malignancies and inherited hematopoietic disorders, is limited by graft-versus-host disease (GVHD) morbidity and mortality (Ferrara et al., 2009). With GVHD, vigorous activation of donor immune cells, most importantly T cells (Korngold and Sprent, 1978), leads to damage of skin, liver, hematopoietic system, and gut. The major sources of immune activation are histocompatibility complex differences between donor and recipient. Combinations of chemotherapy and radiation also contribute, as damage to the intestinal epithelium results in systemic exposure to microbial products normally sequestered in the intestinal lumen (Ferrara et al., 2009). The impact of the microbiota on GVHD is known to be significant. Studies in mice have shown reduction of GVHD with gut-decontaminating antibiotics (van Bekkum et al., 1974) and transplantation in germ-free conditions (Jones et al., 1971). This led to efforts to eliminate bacterial colonization in allogenic BMT patients, combining gut decontamination with a near-sterile environment (Storb et al., 1983). Initial reports were promising, but subsequent studies could not confirm a benefit (Petersen et al., 1987; Passweg et al., 1998; Russell et al., 2000). Other approaches include targeting anaerobic bacteria (Beelen et al., 1999) and introducing potentially beneficial bacteria (Gerbitz et al., 2004), with some reduction of GVHD. These initial studies, however, have been few in number, and no consensus exists between BMT centers regarding how to target the flora. Until recently, a reliance on microbiological culture techniques to characterize flora composition limited these studies. Culture-independent techniques such as ribosomal RNA (rRNA) gene sequencing have demonstrated that a large majority of the estimated 500–1,000 bacterial species present in the human intestinal tract are not detected by culture techniques (Manson et al., 2008). In this study, we readdress the relationship between GVHD and the microbiota in murine and human allogenic BMT recipients. RESULTS AND DISCUSSION Studies of allogeneic BMT using mouse models have characterized exaggerated inflammatory mechanisms that lead to acute GVHD in target organs, including the intestine (Reddy and Ferrara, 2008). MHC-disparate donor/host combinations, including those used in this study, typically result in robust GVHD with full penetrance and rapid kinetics (Schroeder and DiPersio, 2011). The B10.BR→B6 model (H2k→H2b) used in most of our experiments is well-established and has been used in the past by us and others (Blazar et al., 1997, 2000; Penack et al., 2009). On day 14, we evaluated histologically for evidence of GVHD and found villous shortening, increased lymphocytic cell infiltration, crypt regeneration, crypt destruction, and epithelial apoptosis. The number of Paneth cells was also decreased, whereas goblet cells appear to be minimally affected (Fig. 1 A). We then quantified copies of 16S rRNA genes to determine bacterial load. After BMT, we noticed an increase in bacterial load in the ileum (Fig. 1 B), but not in the cecum (unpublished data). This occurred both in the absence and presence of GVHD, suggesting that bacterial expansion may result from reduced host-defense mechanisms in the post-BMT setting. Indeed, we found that levels of IgA in the ileal lumen were decreased after BMT, regardless of GVHD (Fig. 1 C). Figure 1. GVHD in mice produces marked changes in the microbiota. (A) B6 mice were lethally irradiated and transplanted with 5 × 106 B10.BR T cell–depleted BM supplemented with or without 1 × 106 splenic T cells. Features of GVHD are indicated on ileal histology sections from day 14, including lymphocytic infiltration (block arrows), crypt regeneration (enlarged crypts and hyperchromasia), and apoptosis (black arrows). Paneth cells are indicated (blue arrows). Bar, 20 µm. Representative images are shown from one of two independent experiments with similar results. Each dot represents an individual mouse, with bars indicating medians. (B) Quantitation of bacterial load of ileal contents on day 14 was performed by quantitative PCR of 16S rRNA gene copies. Results of a single experiment are shown. (C) Quantification of IgA levels in ileal contents on day 14 was performed by ELISA. Results of a single experiment are shown. (D) Comparison of representation by Unclassified Firmicutes, Barnesiella, and unclassified Porphyromonadaceae from ileal samples. Combined results from three experiments are shown. (E) Diversity of ileal floras from mice with GVHD was determined by the Shannon index. Combined results from two experiments are shown. (F) Principal coordinate analysis of unweighted UniFrac, of ileal floras from B6 mice transplanted with syngeneic (syn) or allogenic (allo) BM with or without T cells. Combined results from three experiments, with data points from each experiment indicated by number. Mice from experiment three were housed individually. (G) Dissimilarity of ileal floras of allo BMT recipient mice without and with GVHD compared with untreated mice by Bray-Curtis index. Combined results of two (Day 7) and 3 (Day 14) experiments are shown. We evaluated for effects on the microbiota by performing 16S rRNA gene sequencing and evaluating microbial diversity, as measured by the Shannon index (Magurran, 2004). Loss of diversity has been found to occur with antibiotic use (Dethlefsen et al., 2008; Ubeda et al., 2010) and increasing age (Woodmansey, 2007), and may predispose mice to disease. Mice undergoing BMT without GVHD showed little change in diversity (unpublished data), but phylogenetic classification of 16S rRNA sequences did show some expansion of unclassified Firmicutes and Barnesiella and mild contraction of unclassified Porphyromonadaceae (Fig. 1 D), demonstrating that radiation does produce some changes in the intestinal flora composition. In contrast, mice with GVHD showed a dramatic loss of bacterial diversity during the first 2 wk after BMT (Fig. 1 E). To quantify changes in the composition of the flora, we used unweighted UniFrac (Lozupone et al., 2006) analyzed by the principal coordinate analysis (PCoA). We found that ileal floras of mice with GVHD were distinct from both those of untreated mice and those of mice after BMT without GVHD (Fig. 1 F). Mice after BMT without GVHD clustered apart from untreated mice inconsistently (one of three experiments); however, comparing the floras using the Bray-Curtis index, we found that GVHD increases dissimilarity from baseline more than BMT alone (Fig. 1 G). We then evaluated for changes in bacterial subpopulations in the setting of GVHD and found large shifts within the phylum Firmicutes, with a dramatic increase in Lactobacillales and decreases in Clostridiales and other Firmicutes in the ileum (Fig. 2 A). Previously, we have shown that the flora from the murine ileum is quite distinct from that of the large intestine, whereas samples within different compartments of the large intestine, including cecum and fresh stool pellets, are similar, although there are some minor changes in representation (Ubeda et al., 2010). Thus, we also evaluated for changes in the cecum with GVHD and found changes similar to those in the ileum, but of lesser magnitude (Fig. 2 B). At the genus level, we found marked ileal expansion of Lactobacillus, the dominant member of Lactobacillales (Fig. 2 C). Housing mice individually from the day of transplant to address the possibility of individual mice influencing the flora of cagemates produced identical results (Fig. 2 C). Within the 16S sequences assigned to the genus Lactobacillus, nearly all had identical sequence homology with Lactobacillus johnsonii, a species found as a commensal in humans (Pridmore et al., 2004) and rodents (Buhnik-Rosenblau et al., 2011), and also in probiotic preparations. Figure 2. GVHD in mice produces marked changes in the microbiota. (A) B6 mice were transplanted with B10.BR donor BM and T cells as in Fig. 1. Comparison of representation by Lactobacillales, Clostridiales, and other Firmicutes from ileal samples. Combined results from three experiments are shown. (B) Comparison of representation by Lactobacillales, Clostridiales, and other Firmicutes from cecal samples. Combined results from two experiments are shown. (C) Bacterial composition at the genus level of ileal flora on day 14 after BMT are depicted with individual mice displayed in each bar. Results of three separate experiments, each displayed in a row, are shown. Additional untransplanted mice were treated with osmotic laxative or DSS starting on day 7 and also individually housed. (D) Mice were transplanted using the strain combinations indicated; mouse vendor was The Jackson Laboratory unless otherwise indicated. Bar graphs show bacterial composition of ileal contents at the genus level for individual mice on day 14. We asked if GVHD-associated changes could be secondary to increased gut motility. We evaluated the effects of an osmotic laxative, as well as enteritis caused by dextran sodium sulfate, and found changes with both agents that were distinct from GVHD (Fig. 2 C). Together, these results suggest that GVHD changes the flora in a unique, reproducible pattern. The flora of mice can vary widely from colony to colony. Our data presented thus far used B6 recipient mice from The Jackson Laboratory; we also performed BMT experiments using additional strains and vendors. With BALB/c host mice from The Jackson Laboratory, we found an abundance of Lactobacillus in mice with GVHD, although the floras of BALB/c mice are dominated by Lactobacillus at baseline (unpublished data). In the CD4-driven MHC II–disparate B6→BM12 model, we also found characteristic expansion of Lactobacillus with GVHD (Fig. 2 D). This indicated that alloreactive CD4 T cells are sufficient and do not require CD8 T cells to produce changes in the flora. Interestingly, in two additional models with B10.BR hosts from The Jackson Laboratory and B6 hosts from Charles River Laboratories, we noted expansion of Enterobacteriales with GVHD (Fig. 2 D). Enterobacteriales from both strains appears to be of the same type, an unclassified Enterobacteriaceae that, in our experience, is rarely detectable in B6 mice from The Jackson Laboratory (3 of 71 mice). Expansion of Enterobacteriaceae has been reported before in Japanese (Eriguchi, Y., S. Takashima, N. Miyake, Y. Nagasaki, N. Shimono, K. Akashi, and T. Teshima. 2010. ASH Annual Meeting Abstracts. Abstr. 244) and German (Heimesaat et al., 2010) mouse colonies. Collectively, these data suggest that Lactobacillales and Enterobacteriales (both capable of surviving in aerobic environments) may populate a niche that expands with GVHD at the expense of obligate anaerobes, including Clostridiales and other Firmicutes. Whether Lactobacillales or Enterobacteriales expand appears to depend on the presence of these organisms in the baseline flora. The potential impact of these expanding populations on GVHD has not been well-described. Treatment of B6 mice with ampicillin, followed by a recovery period, results in loss of Lactobacillus from the flora, with expansion of other commensal bacteria (Ubeda et al., 2010) such as Blautia (order Clostridiales; Fig. 3 A). We cultured the predominant L. johnsonii endogenous to B6 mice from The Jackson Laboratory and found that reintroduction after ampicillin treatment restores representation (Fig. 3 A). We then used ampicillin and L. johnsonii reintroduction as tools to test if expansion of Lactobacillales with GVHD could have clinical repercussions. Surprisingly, upon development of GVHD, mice treated with ampicillin before BMT showed loss of Blautia and emergence of Enterococcus (order Lactobacillales; Fig. 3 A). Mice that received L. johnsonii reintroduction after ampicillin showed domination with L. johnsonii and no expansion of Enterococcus (Fig. 3 A). We found similar results in the B6→BM12 model, though BM12 mice after ampicillin treatment demonstrated expansion of both Enterococcus and Enterobacteriaceae with GVHD (Fig. 3 B). BM12 mice that received L. johnsonii reintroduction after ampicillin also showed domination with L. johnsonii and no expansion of Enterococcus or Enterobacteriaceae. This occurred even when using monoclonal BM12-specific donor T cells from TCR transgenic ABM (Sayegh et al., 2003) RAG-1 deficient mice, suggesting that a broad alloreactive T cell repertoire is not required to produce changes in the microbiota with GVHD. Figure 3. Composition of intestinal flora can impact on severity of intestinal GVHD. (A) Schematic of treatment: B6 mice received ampicillin for 1 wk, followed by a 2-wk recovery period with unmodified drinking water; some were gavaged every 2 d with L. johnsonii (Lacto) of B6 flora origin during recovery, followed by harvest or BMT. Ileal contents were evaluated on days 0 (no BMT) and 14 after BMT. Bar graphs show bacterial composition of ileal contents at the genus level for individual mice. (B) Similar to as in A, BM12 mice received ampicillin followed by recovery; some also received L. johnsonii reintroduction. GVHD was induced upon transplantation with BM and either wild-type CD4 T cells (500K) or ABM RAG1 KO TCR transgenic CD4 T cells (100K). (C) B6 mice were treated with ampicillin, and then were or were not gavaged with L. johnsonii and transplanted with B10.BR BM and T cells. (top) Survival data combined from two experiments with similar results. (bottom) Pathological scores of GVHD target organs on day +21. We then evaluated effects of flora manipulation on GVHD severity, focusing on the B10.BR→B6 model. Notably, ampicillin treatment before BMT resulted in worsened GVHD survival. Histologically, these mice had evidence for increased GVHD pathology in the small and large intestines, including epithelial damage and increased inflammation. Remarkably, L. johnsonii reintroduction prevented increased GVHD lethality and pathology (Fig. 3 C). Enterococcus has not been described as a potential contributor to gut GVHD, though enterococcal bacteremia occurs often in patients with GVHD (Dubberke et al., 2006). In mouse models, Enterococcus can contribute to gut inflammation by compromising epithelial barrier integrity (Steck et al., 2011) and stimulating TNF production from macrophages (Kim et al., 2006). Thus, one mechanism by which L. johnsonii may reduce GVHD severity could be prevention of Enterococcus expansion which may exacerbate GVHD-associated intestinal damage and inflammation. We then studied the relationship between the flora and GVHD in humans. We collected weekly stool samples from allogenic BMT patients during transplant hospitalization at our center. Of 9 patients who developed gut GVHD during hospitalization, 8 developed symptoms early, with GVHD onset clustering between days 18 and 21; these 8 were selected for our GVHD cohort. 18 additional patients provided weekly samples through day 21; of these, 10 met our prospective eligibility for inclusion in our non-GVHD cohort, with survival to at least day 30 and absence of GVHD in any target organ through day 100. Clinical parameters for included patients are summarized in Fig. 4 A. Importantly, non-GVHD and GVHD patients had similar exposures to antibiotics during the period of stool collection. Figure 4. GVHD produces marked changes in the microbiota of humans, and the microbiota may affect risk of developing GVHD. (A) Summary of clinical parameters of non-GVHD and GVHD patients. (B) Flora diversity, by Shannon index, of stool samples after BMT. Individual measurements of diversity are displayed, as well as moving averages and P values calculated for 10-d intervals. (C) Contribution of bacterial populations in samples during two time periods, days 0 to 13 and 14 to 21 after BMT. (D) Microbial chaos of stool samples by mean Bray-Curtis time index from pre-BMT to day 13 after BMT. We first examined the effects of GVHD on flora diversity. We found that before GVHD, patients had flora diversity similar to controls but lost diversity over time, particularly after GVHD onset (Fig. 4 B). Thus, GVHD is associated with loss of flora diversity in humans, similar to in mice. We then looked for bacterial populations that changed with the onset of GVHD. Interestingly, we discovered increases in Lactobacillales and decreases in Clostridiales, a pattern identical to our findings in mice. Other populations, as well as classifications at the family or genus level, were otherwise not significantly changed (unpublished data). Importantly, we did not identify these shifts in non-GVHD patients (Fig. 4 C), suggesting that these flora changes were indeed a result of GVHD rather than BMT or antibiotic exposure. Our sample size did not identify specific populations as potential risk factors for subsequent GVHD. Patients who later developed GVHD, however, did have significantly greater microbial chaos early after BMT (before our observed GVHD-associated changes), which we quantified using the Bray-Curtis dissimilarity index (Magurran, 2004) over time (Fig. 4 D). This suggests that large fluctuations in the microbiota early on may lead to an increased risk of GVHD. In conclusion, our findings demonstrate the influence of inflammation on the structure of the intestinal microbiota after allogenic BMT in both mice and humans. The flora, in turn, can modulate severity of intestinal inflammation. Our mouse experiments indicate that antibiotic exposure before BMT, which occurs commonly in patients with hematologic malignancies, may be a risk factor for subsequent intestinal GVHD. This may be remedied with targeted flora reintroduction to potentially reduce the severity of gut GVHD. MATERIALS AND METHODS Mouse BMT experiments. All mouse procedures were performed in accordance with institutional protocol guidelines at Memorial Sloan-Kettering Cancer Center (MSKCC). Mice were maintained according to National Institutes of Health Animal Care guidelines, under protocols approved by the MSKCC Institutional Animal Care Committee describing experiments specific to this study. Mouse BMT experiments were performed as previously described (Penack et al., 2010). Mice received 11 Gy divided in 2 split doses 3–4 h apart. All BMT experiments were performed at Memorial Sloan-Kettering with the exception of the BALB/c into B6/CR experiment in Fig. 2 D, which was performed at University of Minnesota. All mice were obtained from The Jackson Laboratory, with the exception of B6 mice from Charles River in Fig. 2 D, and ABM mice (Sayegh et al., 2003) in Fig. 3 B, which were provided by M. Sayegh (Brigham and Women’s Hospital and Children’s Hospital Boston, Boston, MA) and had been backcrossed onto a B6 background for at least 20 generations, and then crossed on a RAG-1–deficient background derived from The Jackson Laboratory, previously backcrossed 10 times to B6 background. Mice were either co-housed three to five mice/cage in all experiments, or were housed individually as indicated in experiment three of Fig. 1 F and Fig. 2 C. GVHD clinical and histological scoring. Mice were monitored daily for survival and weekly for GVHD clinical scores (Cooke et al., 1996). Small intestine, large intestine, and liver samples were evaluated histologically for evidence of GVHD and scored as previously described (Hill et al., 1997). Laxative and dextran sodium sulfate (DSS) treatments. Mice were treated with drinking water containing osmotic laxative (60 g/l polyethylene glycol 3350, 1.46 g/l NaCl, 0.745 g/l KCl, 1.68 g/l NaHCO3, and 5.698 g/l Na2SO4) or DSS 3.5% for 7 d. A 7-d course of treatment was selected to better compare with GVHD-induced intestinal changes, which first requires alloactivation and expansion of donor T cells. Ampicillin treatment, Lactobacillus isolation, and reintroduction. Mice were given 1 g/l ampicillin in their drinking water during 7 d, followed by a recovery period with normal drinking water for 14 d. The dominant Lactobacillus strain from the small intestine of B6 mice (The Jackson Laboratory) was isolated by plating contents under anaerobic conditions on plates with Lactobacilli MRS agar (BD). 16S rRNA was sequenced and classified using the ribosomal RDP classifier, and confirmed using MOTHUR to be identical to the operational taxonomic unit (OTU) most predominant in B6 mice (The Jackson Laboratory). 1 d after stopping ampicillin treatment, 108 CFUs of the isolated Lactobacillus strain were given to mice by oral gavage every other day during the 14-d recovery period. Patient selection. We collected stool samples on a weekly basis from allogenic BMT patients from 8/29/09 to 5/24/11. Patients were identified that developed upper gut GVHD symptoms (nausea, vomiting, and loss of appetite) or lower gut GVHD symptoms (abdominal discomfort and diarrhea). 8 had onset of symptoms at a similar time point, between days 18–21, and were selected for our GVHD cohort; the ninth patient had GVHD onset on day 27. 7 underwent confirmatory biopsy; 1 could not be biopsied because of thrombocytopenia. Treatment for GVHD began on days ranging from 26 to 48, and thus the majority of samples that were analyzed were collected before initiation of corticosteroids. Six had symptoms of upper intestinal GVHD and were all treated with the oral corticosteroid budesonide; two also had symptoms of lower intestinal GVHD and were treated with intravenous methylprednisolone. Antibiotic exposures were tabulated from day −7 to 21 after BMT. Standard antibiotic guidelines were followed, including prophylaxis with intravenous vancomycin starting at day −2, and empirical treatment of neutropenic fever with piperacillin/tazobactam, or in patients with allergies, cefepime, or aztreonam. The study protocol was approved by the Memorial Sloan-Kettering Cancer Center Institutional Review Board; informed consent was obtained from all subjects before collection procedures. Sample collection and DNA extraction. Stool samples from patients were stored at 4°C for <24 h before freezing at −80°C. Ileal and cecal samples from mice were frozen at −80°C. DNA was extracted using one of the two methods, which give similar results. In Fig. 1, Fig. 2 (A–C), Fig. 3, and Fig. 4, DNA was extracted using a phenol-chloroform extraction technique (Ubeda et al., 2010). In Fig. 2 D, DNA was extracted from samples using Power Soil DNA isolation kit (MO BIO Laboratories). Quantification of gut flora bacterial density. Gut flora bacterial density was quantified as previously described (Ubeda et al., 2010). IgA quantification. Ileum contents were resuspended in 1 ml of a 3:1 mixture of PBS/0.1 M EDTA containing soybean trypsin inhibitor (type II-S; Sigma-Aldrich) at a concentration of 0.1 mg/ml. The mixture was centrifuged at 12.000 rpm for 10 min, and the supernatant was collected for the assay. Plates were coated with 100 µl/well of rat anti–mouse IgA (SouthernBiotech) at a dilution of 1:1,000 in 50 mM carbonate buffer, pH 9.6. After blocking and washing of plates, 100 µl/well serial dilutions of the previously prepared mouse intestinal samples were added and plates were incubated overnight at room temperature. Bounded antibody was detected by incubating plates at 37°C for 1 h with goat anti–mouse IgA-HRP conjugate at a dilution of 1:1,000 in PBS-T-0.1% BSA. Plates were developed with 2, 2′-azino-bis (3-ethylbenzthiazoline-6-sulfonic acid) (Sigma-Aldrich) and 0.03% H2O2 (Sigma-Aldrich), and optical density was determined using a Vmax microplate reader (Molecular Devices) at 405 nm kinetically for 20 min at 14-s intervals. Total ileum content IgA was calculated using a mouse IgA standard (Kappa TEPC 15; Sigma-Aldrich). 16S rRNA gene amplification, 454 pyrosequencing. For each sample, 3 replicate 25-µl PCRs were performed. Each PCR contained 50 ng of purified DNA, 0.2 mM dNTPs, 1.5 mM MgCl2, 1.25 units of Platinum Taq DNA polymerase, 2.5 µl of 10× PCR buffer, and 0.2 µM of each primer designed to amplify the V1-V2 (Ubeda et al., 2010; Fig. 1 and Fig. 2) or V1-V3 (Fig. 3 and Fig. 4) 16S rRNA variable regions, as described in the Human Microbiome Project Provisional 16S 454 Protocol (http://www.hmpdacc.org/tools_protocols/tools_protocols.php). The cycling conditions used were: 94°C for 3 min, followed by 25 cycles (cecum and fecal samples) or 28 cycles (ileum samples) of 94°C for 30 s, 52°C (V1-V2) or 56°C (V1-V3) for 30 s, and 72°C for 1 min. Replicate PCRs were pooled and amplicons were purified using the QIAquick PCR Purification kit (QIAGEN). PCR products were sequenced on a 454 GS FLX or 454 GS FLX Titanium platform following the recommended procedures (Roche). Sequence analysis. Sequence data were compiled and processed using MOTHUR (Schloss et al., 2009). Sequences were aligned to the 16S rRNA gene, using as a template the SILVA reference alignment and the Needleman-Wunsch algorithm with the default scoring options. Potentially chimeric sequences were removed using the ChimeraSlayer program. To minimize the effect of pyrosequencing errors in overestimating microbial diversity, rare abundance sequences that differ in 1 or 2 nt from a high abundant sequence were merged to the high abundant sequence using the pre.cluster option in MOTHUR. Sequences were grouped into OTUs using the average neighbor algorithm. Sequences with distance-based similarity of 97% or greater were assigned to the same OTU. Mouse samples were processed and sequenced as individual experiments, and resulting sequences were analyzed together with all other samples within each figure panel. Human samples were processed and sequenced in batches, and resulting sequences were analyzed together. Sequences from all experiments have been deposited in the Sequence Read Archive of National Center for Biotechnology Information, submission number SRA049925. Determining diversity, phylogenetic classification, dissimilarity, microbial chaos, and UniFrac PCoA. OTU-based microbial diversity was estimated by calculating the Shannon diversity index (Magurran, 2004) using MOTHUR. Phylogenetic classification was performed for each sequence, using the Bayesian classifier algorithm described by Wang et al. (2007) with the bootstrap cutoff at 60%. A phylogenetic tree was inferred using clearcut on the 16S sequence alignment generated by MOTHUR. Microbial chaos was quantified by mean Bray-Curtis time index, calculated as follows: Bray-Curtis dissimilarity index (Magurran, 2004) between temporally adjacent samples was quantified using MOTHUR and divided by the length of the time interval (in days) between samples, starting with the last sample obtained before the transplant and all samples obtained until day 13. Unweighted UniFrac was run using the resulting tree (Lozupone et al., 2006). PCoA was performed on the resulting matrix of distances between each pair of samples. Statistical comparisons. Shannon diversity index in Fig. 4 B for 10-d intervals compared using unpaired two-sided Student’s t tests with a more stringent cut-off of 0.0125 given multiple comparisons, by the Bonferroni correction for 4 time periods of independent comparisons. Comparisons of bacterial populations in Fig. 3 C using paired two-sided Wilcoxon matched pairs test for individual patients. In Fig. 4 C, Change in Clostridiales was compared using a two-sided Student’s t test, with normality confirmed by D’Agostino and Pearson omnibus test with α = 0.05. All other comparisons were done using two-sided Mann-Whitney tests.
Author and article information
Contributors
Wolf-Hagen Schunck: Role: Editor
Journal
Journal ID (nlm-ta): PLoS One
Journal ID (iso-abbrev): PLoS ONE
Journal ID (publisher-id): plos
Journal ID (pmc): plosone
Title:
PLoS ONE
Publisher:
Public Library of Science
(San Francisco, USA
)
ISSN
(Electronic):
1932-6203
Publication date Collection: 2014
Publication date
(Electronic):
29
January
2014
Volume: 9
Issue: 1
Electronic Location Identifier: e87181
Affiliations
Author notes
* E-mail:
mylesi@
123456niaid.nih.gov
Article
Publisher ID: PONE-D-13-43116DOI: 10.1371/journal.pone.0087181
PMC ID: 3906117
PubMed ID: 24489864
SO-VID: 4fce0ac0-d4cf-439d-9da5-33071511ea8b
Copyright statement:
Copyright @
2014
License:
This is an open-access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.
History
Date
received
: 18
October
2013
Date
accepted
: 20
December
2013
Page count
Pages: 7
Funding
The funders had no role in study design, data collection and analysis, decision to
publish, or preparation of the manuscript. Also the funding was entirely from National
Institutes of Health and the Office of Dietary Supplements. There are no current external
funding sources for this study.
Categories
Subject:
Research Article
Subject:
Biology
Subject:
Biochemistry
Subject:
Lipids
Subject:
Fatty Acids
Subject:
Developmental Biology
Subject:
Genomic Imprinting
Subject:
Microbial Growth and Development
Subject:
Immunology
Subject:
Allergy and Hypersensitivity
Subject:
Genetics of the Immune System
Subject:
Immune Cells
Subject:
Immune Response
Subject:
Immune System
Subject:
Immunomodulation
Subject:
Immunopathology
Subject:
Medicine
Subject:
Infectious Diseases
Subject:
Bacterial Diseases
Subject:
Escherichia Coli
Subject:
Staphylococcal Infection
ScienceOpen disciplines: Uncategorized
Data availability:
ScienceOpen disciplines: Uncategorized