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Which mammals cannot synthesize taurine?

Which mammals cannot synthesize taurine?



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It is fairly common knowledge that domesticated cats cannot synthesize the compound taurine. Other mammals seem to be able to synthesize taurine from cysteine [source]. Are there other mammals that lack the ability to synthesize taurine?


It is a misconception that cats cannot synthesize taurine.

Cats can synthesize taurine, just like other mammals, but not enough of it to make up for an entirely taurine-deficient diet. Cats (and other mammalian carnivores) would have consumed a taurine-rich diet in the ancestral environment. It is only when they are fed vegetable/fruit/grain-derived foods that they show symptoms of taurine deficiency--because those foods have low quantities of taurine and other taurine precursors.

Other domesticated or captive carnivorous mammal species are at risk of taurine-deficiency if they are fed vegetable-derived foods, though members of the cat family (Felidae) are particularly susceptible.


Primary reference: This 2003 paper about taurine concentrations in animal feed, especially the 'Discussion' section.

Taurine is an essential nutrient of cats because the rate of taurine synthesis from its dietary sulphur amino acid precursors, cysteine and methionine, is much less than the extent of loss through faecal bile acids and urine (Knopf et al., 1978). From an evolutionary standpoint, taurine was plentiful in the diet of a true carnivore, as high concentrations of taurine are found in muscle tissue. However, as most domesticated felines normally do not consume living prey, they are at risk to become taurine-deficient if not adequately supplied in the diet.


Physiological role of taurine – from organism to organelle

Correspondence: I. H. Lambert, Dr. Scient., Department of Biology, University of Copenhagen, The August Krogh Building, Universitetsparken 13, DK-2100 Copenhagen Ø, Denmark.

Section of Genomics and Molecular Biomedicine, Department of Biology, University of Copenhagen, Copenhagen, Denmark

Cellular and Metabolic Research Section, Department of Biomedical Sciences, Panum Institute, University of Copenhagen, Copenhagen N, Denmark

Section of Genomics and Molecular Biomedicine, Department of Biology, University of Copenhagen, Copenhagen, Denmark

Cellular and Metabolic Research Section, Department of Biomedical Sciences, Panum Institute, University of Copenhagen, Copenhagen N, Denmark

Section of Cellular and Developmental Biology, Department of Biology, University of Copenhagen, Copenhagen Ø, Denmark

Correspondence: I. H. Lambert, Dr. Scient., Department of Biology, University of Copenhagen, The August Krogh Building, Universitetsparken 13, DK-2100 Copenhagen Ø, Denmark.

Section of Genomics and Molecular Biomedicine, Department of Biology, University of Copenhagen, Copenhagen, Denmark

Cellular and Metabolic Research Section, Department of Biomedical Sciences, Panum Institute, University of Copenhagen, Copenhagen N, Denmark

Section of Genomics and Molecular Biomedicine, Department of Biology, University of Copenhagen, Copenhagen, Denmark

Cellular and Metabolic Research Section, Department of Biomedical Sciences, Panum Institute, University of Copenhagen, Copenhagen N, Denmark

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Abstract

Taurine is often referred to as a semi-essential amino acid as newborn mammals have a limited ability to synthesize taurine and have to rely on dietary supply. Taurine is not thought to be incorporated into proteins as no aminoacyl tRNA synthetase has yet been identified and is not oxidized in mammalian cells. However, taurine contributes significantly to the cellular pool of organic osmolytes and has accordingly been acknowledged for its role in cell volume restoration following osmotic perturbation. This review describes taurine homeostasis in cells and organelles with emphasis on taurine biophysics/membrane dynamics, regulation of transport proteins involved in active taurine uptake and passive taurine release as well as physiological processes, for example, development, lung function, mitochondrial function, antioxidative defence and apoptosis which seem to be affected by a shift in the expression of the taurine transporters and/or the cellular taurine content.


Contents

The major pathway for mammalian taurine synthesis occurs in the liver via the cysteine sulfinic acid pathway. In this pathway, the sulfhydryl group of cysteine is first oxidized to cysteine sulfinic acid by the enzyme cysteine dioxygenase. Cysteine sulfinic acid, in turn, is decarboxylated by sulfinoalanine decarboxylase to form hypotaurine. It is unclear whether hypotaurine is then spontaneously or enzymatically oxidized to yield taurine.

Taurine in the pharmaceutical and lab setting is synthesized through a combination of cysteine, methionine and vitamin E. It is naturally produced in testicles of many mammals. Urban legends surrounding the source of taurine have included bull urine extract and bull semen. While it's true that taurine is found in both sources, it is not the source of taurine in the pharmaceutical or food industry. And while taurine is sometimes extracted from the intestines of cattle, many food industry sources, including the popular energy drink Red Bull, Η] make efforts to use synthesized sources that are vegetarian friendly.


Abstract

Cysteine oxygenase (CDO) is a mononuclear nonhemoglobin enzyme that catalyzes the production of taurine through the cysteine (Cys) pathway and plays a key role in the biosynthesis of taurine in mammals. However, the function of CDOs in bony fish remains poorly understood. In this study, we cloned CDO genes (CaCDO1 and CaCDO2) from Carassius auratus. The cDNA sequences of both CaCDO1 and CaCDO2 encoded putative proteins with 201 amino acids, which included structural features typical of the CDO protein family. Multiple sequence alignment and phylogenetic analysis showed that CaCDO1 and CaCDO2 shared high sequence identities and similarities with C. carpio homologs. Quantitative real-time polymerase chain reaction (qRT-PCR) results revealed that CaCDO1 and CaCDO2 were both broadly expressed in all selected tissues and developmental stages in C. auratus but had differing mRNA levels. In addition, compared to those of the taurine-free group, the in vivo mRNA expression levels of both CaCDO1 and CaCDO2 significantly decreased with increasing dietary taurine levels from 1.0 to 9.0 g/kg. Furthermore, in vitro taurine treatments showed similar inhibitory effects on the expression of CaCDO1 and CaCDO2 in the intestines of C. auratus. Our results also showed that the mRNA expression of CaCDO2 in the intestines was higher than that of CaCDO1 in response to in vivo and in vitro taurine supplementation. Overall, these data may provide new insights into the regulation of fish CDO expression and provide valuable knowledge for improving dietary formulas in aquaculture.


Vitamin B12–dependent taurine synthesis regulates growth and bone mass

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

Find articles by Roman-Garcia, P. in: JCI | PubMed | Google Scholar

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

Find articles by Quiros-Gonzalez, I. in: JCI | PubMed | Google Scholar

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

Find articles by Mottram, L. in: JCI | PubMed | Google Scholar

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

Find articles by Wangwiwatsin, A. in: JCI | PubMed | Google Scholar

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

Find articles by Wilkinson, D. in: JCI | PubMed | Google Scholar

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

Find articles by Santhanam, B. in: JCI | PubMed | Google Scholar

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

Find articles by Vassiliou, G. in: JCI | PubMed | Google Scholar

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

Find articles by Velagapudi, V. in: JCI | PubMed | Google Scholar

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

1 Systems Biology of Bone Laboratory, 2 Sanger Mouse Genetics Project, Department of Mouse and Zebrafish Genetics, 3 Department of Pathogen Genetics, and 4 Department of Human Genetics, Wellcome Trust Sanger Institute, Cambridge, United Kingdom. 5 Instrument Core Facility, University of Aberdeen, Foresterhill, Aberdeen, United Kingdom. 6 Laboratory of Molecular Biology, Medical Research Council, Cambridge, United Kingdom. 7 Pediatrics and Pediatric Hematology, Kocaeli University Hospital, Kocaeli, Turkey. 8 Metabolomics Unit, Institute for Molecular Medicine Finland FIMM, Helsinki, Finland.

Address correspondence to: Vijay K. Yadav, The Morgan Building (N235), Wellcome Trust Sanger Institute, Cambridge CB10 1SA, United Kingdom. Phone: 44.01223.496948 Fax: 44.01223.496826 E-mail: [email protected]

Authorship note: Pablo Roman-Garcia, Isabel Quiros-Gonzalez, and Lynda Mottram contributed equally to this work.

Related video:

Maternal B12 status influences offspring bone mass

The maternal environment not only affects in utero development, but also can dramatically influence postnatal phenotypes. In this episode, Vijay Yadav, Isabel Quiros-Gonzalez, and Liesbet Lieben discuss their use of a murine genetic model to evaluate the effects of maternal vitamin B12 deficiency on offspring bone formation. Pups from B12-deficient mothers exhibited growth retardation and reduced bone mass due to a loss of taurine synthesis in the liver. Furthermore, administration of taurine to these offspring enhanced bone formation, ameliorating growth defects. The results from this study suggest that B12/taurine supplementation may be a potential therapeutic strategy to increase bone mass.

Both maternal and offspring-derived factors contribute to lifelong growth and bone mass accrual, although the specific role of maternal deficiencies in the growth and bone mass of offspring is poorly understood. In the present study, we have shown that vitamin B12 (B12) deficiency in a murine genetic model results in severe postweaning growth retardation and osteoporosis, and the severity and time of onset of this phenotype in the offspring depends on the maternal genotype. Using integrated physiological and metabolomic analysis, we determined that B12 deficiency in the offspring decreases liver taurine production and associates with abrogation of a growth hormone/insulin-like growth factor 1 (GH/IGF1) axis. Taurine increased GH-dependent IGF1 synthesis in the liver, which subsequently enhanced osteoblast function, and in B12-deficient offspring, oral administration of taurine rescued their growth retardation and osteoporosis phenotypes. These results identify B12 as an essential vitamin that positively regulates postweaning growth and bone formation through taurine synthesis and suggests potential therapies to increase bone mass.

The maternal environment plays a fundamental role in the development of the fetus in utero and during postnatal life ( 1 , 2 ). Factors derived from the mother and transported through the placenta, including nutrients and hormones, regulate many physiological processes in the fetus and thus greatly influence late-life health of the offspring ( 3 ). During late gestation and the early postnatal period, there is an exponential deposition of bone in animals, known as bone accrual, that determines the peak bone mass achieved in the adult skeleton ( 4 ). The process that regulates bone accrual, also known as remodeling, consists of 2 phases: resorption of preexisting mineralized bone matrix by the osteoclast, followed by de novo bone formation by the osteoblast ( 5 – 9 ). Despite the importance of the maternal environment in this process, the mechanisms through which maternally derived factors regulate neonatal growth and bone mass are still poorly understood.

In humans, vitamin B12 (B12) deficiency is associated with growth retardation, reduced serum osteocalcin levels, lower bone mineral density, and increased bone fracture risk, yet the underlying mechanisms remain unclear ( 10 – 16 ). B12 is an essential water-soluble vitamin that regulates a multitude of cellular processes in vertebrates ( 17 ). In cells, B12 derivatives function as cofactors for only 2 known enzymes, methionine synthase (MTR) and methylmalonyl-CoA mutase (MUT), and through them affect a variety of downstream metabolic pathways, such as Kreb’s cycle, amino acid synthesis, and DNA and histone methylation ( 18 ). Mammals can recycle B12 to maintain cellular processes dependent on B12 ( 19 ). Absorption of dietary B12 requires gastric intrinsic factor (Gif), a stomach-specific protein that is essential for the absorption of B12 from the gut lumen into the bloodstream ( 17 ), which is then stored in the liver. A decrease in the production of functional Gif protein therefore causes B12 deficiency.

Activation of the hypothalamic/pituitary growth hormone (GH) axis during the early postweaning period in the offspring determines the longitudinal and cross-sectional expansion of the skeleton and other organs in vertebrates ( 20 – 22 ). Perturbations in GH action during early independent life of the offspring result in multiple abnormalities that manifest in skeleton and other organs ( 23 – 26 ). Over the last 20 years, mouse and human molecular genetic studies have identified numerous components in the GH axis that regulate the peripubertal growth spurt ( 23 , 27 ). However, factors regulated by GH in the liver (by which it mediates its actions) are only now beginning to be understood.

Taurine, a sulfur-containing amino acid, is synthesized primarily by liver in the periphery, and after secretion in the circulation, it is accumulated in different tissues of the body ( 28 ). Although taurine is not incorporated in proteins, it is a (semi)essential amino acid for mammals that affects growth and metabolism ( 29 ). Consistent with the critical role of taurine in regulation of growth, its deficiency is often associated with prenatal and postnatal growth retardation ( 29 ). Despite the importance of taurine in regulating various biological functions, its interaction with the GH axis in the regulation of growth and bone metabolism remains undefined.

In the present study, we created a mouse genetic model of B12 deficiency by deleting the gene essential for B12 absorption from the gut, Gif, to understand the importance of maternally and offspring-derived B12 in the regulation of growth and bone mass homeostasis. Through mouse genetic and pharmacological assays using B12-deficient offspring, we showed that maternally derived B12 is an essential nutrient that regulates taurine production in the offspring to regulate growth and bone mass. Importantly, daily oral administration of taurine in B12-deficient offspring was sufficient to prevent their growth defect and osteoporosis through normalization of the GH/IGF1 axis. These results identify B12 as an essential vitamin that regulates growth and bone mass and identify novel avenues to treat bone diseases associated with low bone formation.

B12 deficiency causes growth retardation and low bone mass. To generate B12-deficient animals, we first crossed Gif +/– female mice with Gif +/– male mice, yielding first-generation Gif +/+ [Gif +/+ (F1)] and Gif –/– (F1) mice. Real-time PCR analysis of Gif expression across different tissues showed that Gif expression was restricted to the stomach of WT mice, and this was abolished in Gif –/– (F1) mice (Figure 1A). Measurement of serum B12 levels in Gif –/– (F1) offspring revealed a >20-fold reduction compared with WT mice (600 ± 35 versus 12,000 ± 110 ng/l), yet these mice still harbored detectable levels of serum B12 (Figure 1B). To further deplete B12 levels in the offspring, we next crossed Gif –/– (F1) females with Gif –/– (F1) males to generate second-generation Gif –/– (F2) mice WT control mice were generated by crossing Gif +/+ (F1) females with Gif +/+ (F1) males. Measurement of serum B12 revealed low levels of <45 ng/l (i.e., below the limit of detection of the assay) in Gif –/– (F2) mice, compared with >11,000 ng/l in WT mice (Figure 1B). These results revealed that lowered B12 levels in Gif –/– (F1) females resulted in inadequate B12 transfer to their progeny during pregnancy, which led to very low levels of B12 in their serum.

B12 deficiency in mice causes growth retardation and low bone mass. (A) Real-time PCR analysis of Gif expression in WT and Gif –/– tissues. (B) Serum B12 levels in WT, Gif –/– (F1), and Gif –/– (F2) mice. (C) BW analysis of WT, Gif –/– (F1) and Gif –/– (F2) mice. (D) Morphological analysis of 8-week-old WT, Gif –/– (F1), and Gif –/– (F2) mice. (E and F) Histological analysis of vertebrae (E) and μCT analysis of long bone (F) of WT, Gif –/– (F1), and Gif –/– (F2) mice. Mineralized bone matrix (black) was stained by von Kossa reagent. BV/TV, bone volume relative to total volume. Ct.Th., cortical thickness. (G) Toluidine blue staining showing reduced osteoblast number on bone surface, with quantification of Ob.N/T.Ar. (H) Photomicrographs showing near-absence of calcein double labeling on the surface of trabecular bone in Gif –/– (F2) mice, with quantification of BFR. (I) Photomicrographs showing TRAP-stained osteoclasts on the bone surface (pink), with quantification of osteoclast surface per bone surface (OcS/BS). # P < 0.05 *P < 0.01. Values are mean ± SEM. n = 8 [WT and Gif –/– (F2)] 9 [Gif –/– (F2)]. Arrowheads on images indicate the location of cell types or parameters measured. Scale bars: 10 mm (D) 1 mm (E and F) 0.1 mm (G) 50 μm (H) 10 μm (H, insets) 0.05 mm (I). See also Supplemental Figure 1.

We used Gif –/– (F1) and Gif –/– (F2) mice to address the effect of altered B12 levels on postnatal growth and bone mass accrual. There was no major difference in the growth of WT, Gif –/– (F1), and Gif –/– (F2) animals until P21 (Figure 1C and Supplemental Figure 1, A and B supplemental material available online with this article doi:10.1172/JCI72606DS1). However, Gif –/– (F2) animals showed growth arrest between P21 and P26, as evidenced by their impaired BW gain and reduced body size compared with WT, Gif –/– (F1), and Gif +/– (F2) animals (Figure 1, C and D, and Supplemental Figure 1C). These results indicated that although Gif –/– (F2) mice have normal early postnatal development, the postweaning growth spurt observed in WT animals is severely compromised in these mice.

Histological and μCT analyses in 8-week-old WT, Gif –/– (F1), and Gif –/– (F2) mice revealed a severe decrease in bone mass in vertebra and long bone in Gif –/– (F2) animals, whereas levels were similar between Gif –/– (F1) and WT mice (Figure 1, E and F). The low bone mass in Gif –/– (F2) mice was caused by a >3-fold decrease in number of osteoblasts per trabecular area (Ob.N/T.Ar.), with a dramatic decrease in bone formation rate (BFR), compared with WT mice (Figure 1, G and H). In contrast to the deleterious consequences of B12 deficiency on osteoblast number and function, osteoclast parameters in Gif –/– (F2) mice were similar to those of WT mice (Figure 1I).

Together, these results obtained in first- and second-generation Gif –/– offspring showed (a) that maternally derived B12 is transferred through at least 2 generations (in the first, it is sufficient to support offspring growth, but further depletion in the second fails to support their postweaning growth spurt) and (b) that B12 deficiency in the offspring decreases bone mass by reducing osteoblast numbers and bone formation without affecting bone resorption.

A single injection of B12 to the mothers prevents growth retardation and osteoporosis in Gif –/– (F2) offspring. To address whether the growth retardation and osteoporosis in Gif –/– (F2) progeny was due to the inability of their mothers to transfer adequate B12, we next gave a single injection of B12 to Gif –/– (F1) mothers. Pregnant WT and Gif –/– (F1) females were given either vehicle or a single s.c. injection of 200 μg cyanocobalamin (CN-B12) at 12.5 days post coitum (dpc), and their offspring [referred to herein as Gif –/– (F2)/VEH and Gif –/– (F2)/B12, respectively] were studied. Growth curve analysis showed that although Gif –/– (F2)/B12 offspring, like Gif –/– (F2)/VEH mice, showed growth retardation from P21 to P26, they rapidly caught up with WT mouse growth between P26 and P31, and were in fact indistinguishable from WT animals at P31 and thereafter (Figure 2A). In addition, bone histology and histomorphometry analysis at 8 weeks of age showed that the low bone mass observed in Gif –/– (F2)/VEH mice was completely prevented, in both vertebra and long bones, in Gif –/– (F2)/B12 mice to levels seen in WT mice, due to the normalization of bone formation parameters (Figure 2, B and C). Growth curve and bone parameters were also normalized in male Gif –/– (F2)/B12 mice to levels seen in WT mice (Supplemental Figure 2, A–I). Thus, a single s.c. injection of CN-B12 to Gif –/– (F1) mothers was sufficient to rescue growth retardation and bone loss in their progeny at 8 weeks of age. Although Gif –/– (F2)/B12 offspring had similar BW and bone mass at 8 weeks of age, they still displayed growth retardation during P21–P26, indicative of a higher requirement for B12 during this period. To test this contention, we next gave a single injection of B12 to the offspring on P11, closer to growth retardation onset at P21. This single injection to the offspring fully rescued their growth parameters up to 24 weeks of age (Supplemental Figure 2J and data not shown), which indicates that the exponential growth observed after weaning in offspring indeed requires a higher amount of B12.

Maternal B12 regulates offspring growth and bone mass and B12 deficiency during aging regulates bone mass independently of body growth. (A) Growth curve analysis of WT/VEH, Gif –/– (F2)/VEH, and Gif –/– (F2)/B12 mice. (B and C) Histological analysis of vertebral bone (B) and μCT analysis of long bone (C) in 8-week-old WT/VEH, Gif –/– (F2)/VEH, and Gif –/– (F2)/B12 mice, with quantification of BV/TV, Ob.N/T.Ar., BFR, OcS/BS, and Ct.Th. (D and E) BW (D) and nose-to-tail length (E) of 48-week-old WT and Gif –/– (F1) mice. (F and G) Histological analysis of vertebral bone (F) and Ct.Th. analysis of long bone (G) in 48-week-old WT and Gif –/– (F1) mice, with quantification of BV/TV, Ob.N/T.Ar., OcS/BS, and Ct.Th. # P < 0.05 *P < 0.01. Values are mean ± SEM. n = 4–6 [WT] 5 [Gif –/– (F2)/VEH], 7 [Gif –/– (F2)/B12] 6 [Gif –/– (F1)]. All mice shown are females. Scale bars: 1 mm (B, C, and F). See also Supplemental Figure 2.

B12 deficiency during aging regulates bone mass independently of body growth. Our observation that the maternally derived pool of B12 in Gif –/– (F1) mice was sufficient to support their postnatal growth and bone mass until adulthood provided us with a model wherein we can use aged F1 mice to dissociate the effect of maternal B12 on bone mass from its effect on growth. We reasoned that because Gif –/– (F1) mice do not have the ability to absorb B12 from their diets and only receive a finite amount of B12 from their mothers, they would eventually deplete this store, become B12 deficient, and develop osteoporosis. Consistent with this model, analysis of 48-week-old Gif –/– (F1) mice revealed normal growth, as evidenced by normal BW and nose-to-tail length, compared with WT mice (Figure 2, D and E). In contrast to their normal BW, 48-week-old Gif –/– (F1) mice displayed a severe decrease in bone mass compared with WT animals in both vertebra and long bones (Figure 2, F and G). This low bone mass was associated with a >2-fold decrease in Ob.N/T.Ar. compared with WT mice, without any changes in osteoclast parameters (Figure 2F). These results demonstrated that B12 deficiency during aging in the offspring affects bone mass independent of body growth.

B12 deficiency causes GH resistance. Given that loss of B12 caused low bone mass and decreased osteoblast numbers in vivo, we considered the possibility that B12 may regulate osteoblast numbers and function by directly acting on these cells to regulate bone mass. To test this contention, we cultured primary osteoblasts in B12-deficient medium, treated them with different doses of B12, and measured their proliferation and differentiation. Reducing the concentration of B12 from 10,000 to 0 nM did not have any significant effect on proliferation or differentiation of osteoblasts (Supplemental Figure 3A). Consistent with these results, primary osteoblasts from B12-deficient mice proliferated and differentiated equally well as WT cells (Supplemental Figure 3A). These in vitro results were surprising, in contrast to the in vivo results of a ≥3-fold decrease in osteoblast numbers and function in B12-deficient animals, which suggests that B12 deficiency likely affects osteoblasts through an endocrine mechanism.

GH, a hormone made in the pituitary gland, appeared to be the likely candidate, given its role in postweaning growth and bone mass accrual ( 23 , 27 , 30 ). We therefore first analyzed serum levels of GH in 8-week-old WT and Gif –/– (F2) mice. Serum GH levels were increased approximately 3-fold in mutants compared with WT mice (Figure 3A). This increase in GH in Gif –/– (F2) animals was associated with increased expression of GH releasing hormone (Ghrh), which regulates pituitary GH production, in the hypothalamus (Figure 3B). These results suggested that B12-deficient animals display GH resistance or insensitivity, at the level of either GH receptor (Ghr) or one of its downstream effectors, in the liver. Analysis of Ghr showed normal expression in the liver and bone in Gif –/– (F2) animals however, levels of serum IGF1 (through which GH mediates many of its peripheral actions) were reduced >4-fold (Figure 3, C and D). The decrease in IGF1 serum levels was associated with a decrease in IGF1 receptor (IGF1R) phosphorylation in the target tissues (Figure 3E).

B12 deficiency causes GH resistance. (A) Serum GH levels in WT (n = 6) and Gif –/– (F2) (n = 5) mice. (B and C) Real-time PCR analysis of Ghrh in the hypothalamus (Hyp) (B), and Ghr expression in liver and bone (C), of WT and Gif –/– (F2) mice (n = 5 per group). (D) Serum IGF1 levels in WT and Gif –/– (F2) mice (n = 7–10). (E and F) Western blot analysis of IGF1R (E) and STAT5 (F) phosphorylation in different WT and Gif –/– (F2) tissues blotting was done on the same blot after stripping the membrane for pIGF1R and pSTAT5, respectively. A representative blot from 3 independent experiments is shown different tissues were run noncontiguously. Relative quantification of pIGF1R and pSTAT5 (normalized to IGF1R and STAT5, respectively) is shown below. (G and H) Real-time PCR analysis of Socs2 (G) and STAT5 target gene (H) expression in WT and Gif –/– (F2) liver (n = 5 per group). (I) Enzymatic reactions of MTR and MUT dependent on the B12-generated cofactors methyl-B12 and adenosyl-B12, respectively. (J) Levels of methionine, succinate, and homocysteine (nmol/g liver tissue) in WT and Gif –/– (F2) mice (n = 5 per group). (K) ChIP analysis of methylated histones, shown relative to control (assigned as 1), in different regions of Igf1 in Gif –/– (F2) liver. P-, promoter E-, exon n.d., not detectable. (L) Survival of WT/VEH, WT/IGF1, Gif –/– (F2)/VEH, and Gif –/– (F2)/IGF1 mice (n = 5 per group). # P < 0.05 *P < 0.01. Values are mean ± SEM. See also Supplemental Figure 3.

Since Igf1 expression is regulated by STAT5 signaling downstream of GH, we next investigated whether activation of STAT5 and its major targets is perturbed in Gif –/– (F2) mice. In Gif –/– (F2) mice, STAT5 phosphorylation was nearly absent in the liver and muscle, 2 major GH target tissues, but not in other tissues (Figure 3F). The decrease in STAT5 phosphorylation was associated with a major increase in expression of Socs2, an inhibitor of STAT5, in Gif –/– (F2) liver (Figure 3G). In addition, mRNA levels of major STAT5 targets in the liver ( 31 ), such as IGF binding protein 1 (Igfbp1), was increased, whereas expression of Igf1, IGF binding protein acid labile subunit (Igfals), solute carrier organic anion transporter family member 1a1 (Slco1a1), major urinary proteins (Mup1Mup3), and hydroxysteroid dehydrogenase 3b5 (Hsd3b5) was decreased (Figure 3H and Supplemental Figure 3B), similar to previous observations in Stat5 –/– mice ( 32 ). This GH resistance was also present in aged Gif –/– (F1) mice, and maternal B12 injections normalized GH/IGF1 levels in Gif –/– (F2) offspring (Supplemental Figure 3, C and D). These results revealed that B12 deficiency results in major abrogation of GH action in the offspring.

We next investigated how B12 deficiency abrogates GH action, using liver tissue as a model. B12 derivatives in mammals act as cofactors for the function of only 2 known enzymes, MTR and MUT (Figure 3I) therefore, we first measured the metabolites downstream of these enzymes to observe the effect of B12 deficiency on their function. Methionine, a product of MTR, was significantly downregulated, and its substrate, homocysteine, was increased, whereas succinate, a downstream product of MUT, was not affected (Figure 3J). Methionine is a precursor for cellular production of S-adenosyl methionine (SAM), which is an essential methyl donor for histone and DNA methylation, 2 epigenetic modifications known to affect basal and ligand-stimulated gene expression levels ( 33 ). Because we observed a major decrease in the expression of liver Igf1, we first tested whether B12 deficiency alters the methylation status of this gene, using histone methylation as a marker. ChIP analysis of histone methylation in WT and Gif –/– (F2) liver tissues revealed a major decrease in histone (H3) methylation at the Igf1 locus (Figure 3K). These results suggested that a major decrease in histone methylation might lead to decreased responsiveness of target tissues to GH, resulting in development of GH resistance.

If the observed decrease in liver IGF1 synthesis alone is necessary and sufficient to cause growth retardation and osteoporosis in Gif –/– (F2) animals, then IGF1 administration to these animals should be able to overcome their growth retardation. To examine this possibility, we treated Gif –/– (F2) animals beginning at P20 with twice-daily injections of recombinant IGF1, which has previously been shown to prevent and/or cure growth abnormalities ( 34 ). To our surprise, IGF1 treatment led to early lethality of Gif –/– (F2) animals (Figure 3L). This extreme response to IGF1 can be explained by the lower basal glucose levels observed in these animals (15.1 ± 1.2 versus 25.5 ± 3.2 nM), likely due to GH resistance, and IGF1 administration further decreased their glycemic state (5.2 ± 1.3 nM at this point, if animals were given a bolus of glucose, reviving them and preventing death), likely resulting in lethality (Figure 3L). Moreover, administration of SAM that normalized DNA methylation could not rescue the growth retardation and bone abnormalities caused by B12 deficiency (data not shown). These results suggest that the suppression and/or activation of another GH mediator in addition to SAM (dependent on B12, either acting independently or regulating IGF1 action) underlies GH resistance in B12-deficient animals.

Identification of taurine as a critical metabolite positively regulated by B12. To identify the mediators underlying the GH resistance observed upon B12 deficiency, we next carried out targeted metabolomic analysis in the liver of Gif –/– (F2) mice and compared this profile with GH-regulated metabolome. Analysis of metabolomic data upon B12 deficiency revealed that most of the metabolites were downregulated and were involved in amino acid, betaine, primary bile acid, protein, and purine metabolism (Figure 4A and Supplemental Figure 4A). Metabolite profiling of GH-treated hepatocytes identified many metabolites that showed similar positive regulation by GH and B12 (Figure 4A and Supplemental Figure 4B) and might underlie the GH resistance observed upon B12 deficiency.

Metabolomics analysis identifies taurine as a critical metabolite that connects B12 deficiency with GH signaling. (A) Supervised hierarchical clustering plot of up- or downregulated metabolites in Gif –/– (F2) liver. Metabolites regulated by GH in hepatocytes are shown in red font. (B) Summary plot for quantitative enrichment analysis. Metabolite sets are ranked according to false discovery rate (FDR) dashed lines show FDR value cutoffs. (C) Metabolome view reflects on the x axis increasing metabolic pathway impact according to the betweenness centrality measure, which shows key nodes in metabolic pathways that have been significantly altered upon B12 deficiency. Colored circles correspond to pathways in B. (D) PLSDA-VIP plot. Metabolites are ranked according to their increasing importance to group separation between WT and Gif –/– (F2) mice. (E) Measurement of taurine and its derivatives in WT/VEH, Gif –/– (F2)/VEH, and Gif –/– (F2)/B12 liver (n = 5 per group). # P < 0.05 *P < 0.01. Values are mean ± SEM. See also Supplemental Figure 4.

Quantitative enrichment analysis on B12-deficient metabolome showed major perturbations in the metabolic pathways associated with taurine and hypotaurine, folate and bile acid biosynthesis, nucleotide and amino acid metabolisms, and protein synthesis (Figure 4B). Pathway analysis identified that the taurine and hypotaurine pathways had a substantial effect on cellular function following B12 deficiency (Figure 4C). To identify which metabolites contribute to the group separation of WT and B12-deficient samples, we performed a supervised multivariate regression technique, partial least squares discriminant analysis (PLSDA). In the PLSDA model, the number of latent variables (LVs) to be used depends on the sum of squares captured by the model (R 2 ), cross-validated R 2 (Q 2 ), and prediction accuracies based on cross-validations, with different numbers of LVs. Variable importance in projection (VIP) is one of the important measures of PLSDA, where it is a weighted sum of squares of the PLS loadings taking into account the amount of explained class variation in each dimension. PLSDA-VIP analysis of the liver metabolomics data identified that taurine had the highest VIP score and could serve as a biomarker for B12-deficient status (Figure 4D). Consistent with the notion that perturbation in taurine metabolism underlies the GH resistance observed upon B12 deficiency, measurement of taurine and its derivatives in Gif –/– (F2)/B12 liver showed complete rescue of these metabolites to the levels seen in WT liver (Figure 4E). Moreover, further suggesting a critical role of taurine in the GH/B12-dependent pathway, taurine deficiency has previously been shown to be associated with GH/IGF1 signaling downregulation in Ames dwarf mice ( 35 ), and methionine metabolism is significantly perturbed in patients with GH deficiency ( 36 ).

Taken together, these integrated metabolomic analysis in WT and Gif –/– (F2) mouse liver identified a hitherto-unanticipated suppression of taurine metabolism in B12 deficiency–mediated GH resistance.

GH regulates taurine synthesis in a STAT5/B12-dependent manner. Thus far, our results provided correlative evidence that B12 deficiency in the liver results in reduced taurine production associated with abrogation of GH signaling. We next generated in vitro models of B12 and STAT5 deficiency in hepatocytes and asked whether GH/STAT5 signaling regulates taurine production and, if so, whether this process is affected by B12 deficiency (Figure 5A). We used 2 strategies to investigate this question. First, to investigate the B12 regulation of GH-dependent taurine synthesis, we used a sequestration approach to ablate B12 action in hepatocytes using overexpression of a transcobalamin 2–oleosin construct (TCOL) in HepG2 cells. Upon expression, TCN2 binds to B12 in the cytoplasm, and oleosin immobilizes this TCN2-B12 complex to the endoplasmic reticulum, thereby chelating any B12 present in the cytosol ( 37 ). To this end, we synthesized TCOL fusion vectors and created stably transfected HepG2 cells with TCOL or an empty vector as a control. Measurement of Oleosin transcript in the cells showed that we successfully overexpressed TCOL (Figure 5B). Measurement of taurine in the transfected cells showed that while GH treatment significantly increased pSTAT5 and taurine production in the vector-transfected cells, this increase in pSTAT5 and taurine was completely abrogated in TCOL-transfected cells (Figure 5, C and D). PLSDA-VIP analysis of the metabolomics data derived from these cells showed that, once again, taurine was the main driver for the separation of the 3 groups (Figure 5E). Importantly, addition of taurine to TCOL-transfected cells fully restored GH responsiveness, as measured by IGF1 expression analysis (Supplemental Figure 5A).

GH regulates taurine synthesis in a STAT5- and B12-dependent manner. (A) Relationship among taurine, STAT5, and B12 in the liver. (B) Experimental regimen used to test B12 involvement in GH regulation of taurine synthesis. Also shown is RT-PCR analysis to detect Oleosin transcript in the cells after transfection with empty vector or TCOL construct. (C) STAT5 phosphorylation upon GH treatment in empty vector– or TCOL-transfected HepG2 cells. Blots were run noncontiguously. (D) Taurine levels upon GH treatment in empty vector– or TCOL-transfected HepG2 cells. (E) PLSDA-VIP scores plot of metabolomics data from hepatocytes after empty or TCOL transfection. (F) Experimental regimen used to test STAT5 involvement in GH regulation of taurine production. Photomicrographs show immunohistochemistry of STAT5 in HepG2 cells transfected with nontargeting (empty) or STAT5 (shSTAT5) shRNA. (G) Relative expression of IGF1 upon GH treatment in empty or STAT5 shRNA–transfected HepG2 cells. (H) Taurine levels upon GH treatment in empty or STAT5 shRNA–transfected HepG2 cells. (I) Real-time PCR analysis of enzymes in the taurine synthesis pathway in empty, TCOL, or STAT5 shRNA–transfected cells treated with vehicle or GH. (J) GH regulation of the taurine synthesis pathway. Red metabolites and genes, upregulated (only those upregulated by GH and that do not respond to GH upon STAT5 shRNA or TCOL transfection) green metabolites and genes, downregulated black metabolites and genes, not altered gray metabolites and genes, not measured. *P < 0.05 # P < 0.01. Values are mean ± SEM. Scale bars: 20 μm (F). See also Supplemental Figure 5.

The observations that B12-deficient animals had a major decrease in STAT5 phosphorylation, and that GH increased taurine production, led us to next investigate whether GH — through STAT5 signaling — regulates taurine production. We stably transfected hepatocytes (HepG2 cells) with shRNA against STAT5 or empty vector, followed by treatment of these cells with either vehicle or GH. Immunohistochemical analysis of STAT5 levels in these cells after transfection revealed that STAT5 expression was successfully abrogated in STAT5 shRNA–transfected compared with mock-transfected cells (Figure 5F). Measurement of IGF1 expression confirmed that we successfully abrogated GH action (Figure 5G). We next measured taurine production in response to GH. GH treatment significantly increased taurine production in the mock-transfected cells, but this increase was completely abrogated in STAT5 shRNA–transfected cells (Figure 5H).

Finally, we questioned which of the enzymes in the taurine synthesis pathway are regulated by GH in a STAT5- and B12-dependent manner. GH treatment of hepatocytes lead to an increase in the expression of gamma-glutamyltransferase 6 (GGT6), cystathionine beta synthase (CBS), cysteine sulfinic acid decarboxylase (CSAD), dimethylglycine dehydrogenase (DMGDH), S-adenosylhomocysteine hydrolase-like 1 (AHCYL1), aldehyde dehydrogenase 7 family, member A1 (ALDH7A1), and glycine N-methyltransferase (GNMT), while it did not affect expression of other enzymes (BHMT, SHMT1, BAAT, and MAT1A) in this pathway (Figure 5I and data not shown). The increase in expression of these enzymes observed upon GH treatment was abrogated following loss of STAT5 (STAT5 shRNA) or with sequestration of B12 in the cells (Figure 5I). Integrated analysis of GH-regulated metabolites and gene expression changes linking methionine to taurine metabolism showed that GH signaling affected taurine biosynthesis pathway at multiple levels (Figure 5J).

Based on these results, we concluded (a) that GH increases taurine production in the hepatocytes in a STAT5- and B12-dependent manner and (b) that STAT5 and B12 regulate taurine synthesis by regulating expression of critical enzymes and metabolites in the taurine synthesis pathway. Together, these results showed that GH signals in the hepatocytes in a STAT5/B12-dependent manner to regulate production of taurine, whose deficiency in B12-deficient animals might underlie GH resistance and osteoporosis.

Oral feeding of taurine to B12-deficient Gif –/– (F2) offspring normalizes the GH/IGF1 axis and prevents their growth retardation and osteoporosis. If the decrease in taurine levels is the cause of GH resistance and low bone mass phenotype in Gif –/– (F2) animals, then administration of taurine to these animals should be able to prevent postweaning growth and bone mass abnormalities. To this end, 16-day-old WT and Gif –/– (F2) females were given either vehicle or 500 mg/kg BW taurine daily (orally) [referred to herein as Gif –/– (F2)/VEH and Gif –/– (F2)/TAU mice, respectively]. Growth curve analysis showed that although Gif –/– (F2)/VEH offspring showed growth retardation from P21 to P26, Gif –/– (F2)/TAU mice grew normally between P21 and P26 and thereafter, and were in fact indistinguishable from WT animals (Figure 6A). This rescue of growth retardation in Gif –/– (F2)/TAU animals was associated with normalization of serum GH and IGF1 levels, which were restored to the levels seen in WT animals (Figure 6, B and C). PLSDA analysis of liver metabolite profiles revealed that Gif –/– (F2)/TAU mice clustered with the WT mice, whereas Gif –/– (F2)/VEH mice were clearly distinct from these 2 groups (Supplemental Figure 6A). This rescue of liver metabolome was also reflected in 2-way clustering performed on the metabolite profiles (Supplemental Figure 6B). Histological and histomorphometry analysis of bone from 8-week-old WT, Gif –/– (F2)/VEH, and Gif –/– (F2)/TAU mice revealed that the low bone mass observed in Gif –/– (F2)/VEH mice was completely rescued in Gif –/– (F2)/TAU mice, due to a major increase in osteoblast number and function, while osteoclast parameters were not affected (Figure 6D and Supplemental Figure 6C). Thus, daily oral administration of taurine to B12-deficient offspring prevented the growth retardation and low bone mass caused by B12 deficiency. These results indicated that taurine is an essential downstream mediator of B12-dependent GH signaling in the regulation of growth and bone metabolism. The observation that rescue of growth and bone mass abnormalities in taurine-treated B12-deficient mice was associated with a major increase in serum levels of IGF1, a major regulator of osteoblast function, led us to next investigate the importance of IGF1 signaling in this process. We performed a correlation analysis between bone mass or osteoblast numbers and serum levels of IGF1 to investigate whether IGF1 signaling contributes to the changes in bone mass in taurine-fed B12-deficient animals. This analysis showed a linear correlation between serum IGF1 levels and bone mass or osteoblast numbers in taurine-fed animals (Figure 6E and Supplemental Figure 6D). Together, these results suggest that IGF1 synthesis from the liver and its action in IGF1 target tissues underlies taurine-mediated rescue of growth retardation and osteoporosis in Gif –/– (F2) mice.

Oral taurine administration prevents growth retardation and osteoporosis in Gif –/– (F2) mice. (A) Growth curve analysis of WT/VEH, Gif –/– (F2)/VEH, and Gif –/– (F2)/TAU mice. (B and C) Serum GH (B) and IGF1 (C) levels in WT/VEH, Gif –/– (F2)/VEH, and Gif –/– (F2)/TAU mice. (D) Histological analysis of vertebra in WT, Gif –/– (F2)/VEH, and Gif –/– (F2)/TAU mice, with quantification of BV/TV, Ob.N/T.Ar., BFR, and OcS/BS. (E) Pearson correlation scatter plot between BV/TV and serum IGF1 levels in taurine-treated mice. *P < 0.05 # P < 0.01. n = 5 per group. Values are mean ± SEM. Scale bar: 500 μm. See also Supplemental Figure 6.

Taurine increases IGF1 synthesis in hepatocytes and its action in osteoblasts to regulate bone mass. We next investigated the mechanisms through which taurine affects liver IGF1 synthesis downstream of GH to regulate bone mass. The taurine-mediated increase in IGF1 synthesis suggested that taurine was able to overcome the consequences of B12 deficiency in the liver. Analysis of metabolite levels in the liver showed that methionine levels in taurine-treated animals were rescued to the levels seen in WT animals (Figure 7A). Because MTR, the principle enzyme that synthesizes methionine in mammals, cannot function in the absence of B12, we looked at alternate pathways of methionine synthesis that are not dependent on B12 as a cofactor and could explain the increase in methionine synthesis in response to taurine. Analysis of protein levels and activity of betaine-homocysteine S-methyltransferase (BHMT), an enzyme that carries out a parallel reaction to synthesize methionine, revealed a major increase in its protein levels in the liver of taurine-fed animals (Figure 7, B and C). Consistent with the increased protein levels of BHMT, its substrates (betaine and homocysteine) were decreased, whereas its products (dimethylglycine and methionine) were increased, in liver (Figure 7, B and D). Bhmt expression was found to be undetectable in bone (Supplemental Figure 7A). ChIP analysis of me2K36, a marker of histone methylation levels, at the Igf1 locus showed that this increase in methionine levels normalized the methylation status of Igf1 in the liver (Supplemental Figure 7B). In contrast to the changes in liver Igf1, methylation status of Igf1 in osteoblasts or its expression in the long bone was not different among WT, Gif –/– (F2)/VEH, and Gif –/– (F2)/TAU mice (Supplemental Figure 7, C and D). Together, these results revealed that taurine-mediated BHMT pathway activation led to the restoration of methylation status and responsiveness of Igf1, and other genes, to circulating GH.

Taurine increases IGF1 synthesis from liver and its action in osteoblasts to regulate bone mass. (AD) Liver samples. (A) Levels of methionine and homocysteine in WT, Gif –/– (F2)/VEH, and Gif –/– (F2)/TAU liver. (B) B12-dependent (MTR) and -independent (BHMT) methionine synthesis pathways. (C) Western blot analysis of BHMT levels in WT, Gif –/– (F2)/VEH, and Gif –/– (F2)/TAU liver. Lanes were run contiguously. U.D., undetectable. (D) Levels of betaine and dimethyl-glycine in liver of WT, Gif –/– (F2)/VEH, and Gif –/– (F2)/TAU mice. (EG) MC3T3-E1 osteoblast cells. Changes in BrdU incorporation (E), IGF1R and ERK phosphorylation (F), and Ccnd1 expression (G) in cells treated for 24 hours with vehicle, OSI906, taurine, or taurine plus OSI906. Lanes in F were run contiguously, and blots were stripped and reprobed with IGF1R or ERK. Relative quantification of pIGF1R and pERK (normalized to IGF1R and ERK, respectively) is shown below. A representative blot from 3 different experiments is shown. (H) Growth curve analysis of WT and Gif –/– (F2)/TAU+OSI906 mice (n = 5 each). (I) Bone mass analysis (BV/TV) in the vertebra of WT and Gif –/– (F2)/TAU+OSI906 mice (n = 5 per group). (J) Gut/liver/bone endocrine axis, illustrating GH/STAT5/B12-dependent changes in serum IGF1 and taurine that regulate osteoblast proliferation and bone mass. *P < 0.05 # P < 0.01. Values are mean ± SEM. Scale bar: 500 μm. See also Supplemental Figure 7.

Next, we investigated which aspects of osteoblast biology are affected by taurine, and whether taurine’s effect is dependent on IGF1 signaling. Taurine treatment (20 mM) specifically increased osteoblast proliferation, but had no effect on the differentiation or mineralization of these cells, and this increase in osteoblast proliferation was blocked by pretreatment with the IGF1R antagonist OSI906 (10 μM) (Figure 7E and Supplemental Figure 7, E–G). Analysis of signaling pathways downstream of taurine in osteoblasts revealed an increase in IGF1R phosphorylation and downstream activation of the ERK pathway, as reflected by increased pERK levels and increased expression of its transcriptional target, Ccnd1, both of which were abrogated by OSI906 pretreatment (Figure 7, F and G). This effect of taurine on IGF1R signaling was also seen in the long bone in vivo (Supplemental Figure 7, H and I). These results showed that taurine stimulates the IGF1R/ERK signaling cascade to increase osteoblast proliferation.

Finally, to investigate the importance of IGF1R in mediating the bone and growth abnormalities in Gif –/– (F2) mice, we treated these mice with either vehicle [Gif –/– (F2)/VEH] or taurine plus OSI906 (50 mg/kg BW/d) [Gif –/– (F2)/TAU+OSI906] from P16 onward. Growth curve and bone histomorphometric analyses showed that Gif –/– (F2)/TAU+OSI906 mice remained growth retarded and osteoporotic (Figure 7, H and I). These results indicated that B12/taurine is an essential mediator of GH action that increases IGF1 synthesis from the liver and its signaling in osteoblasts to increase osteoblast proliferation and bone mass while it increases GH signaling in other tissues, such as cartilage, which is dependent on GH directly to increase longitudinal growth (Supplemental Figure 7J).

Evidence for the B12/taurine/bone pathway in children and aged patients with B12 deficiency. Serum samples from children born of nutritionally B12-deficient mothers showed a significant decrease in their serum B12, taurine, and osteocalcin (as a marker of bone formation) levels compared with age-matched controls (Supplemental Figure 8, A and B). Correlation analysis using all samples showed a significant positive correlation between variation in serum B12 and taurine (R = 0.877, P = 0.00018), B12 and osteocalcin (R = 0.830, P = 0.00083), and taurine and osteocalcin (R = 0.742, P = 0.0057) levels (Figure 8, A–C).

B12 status correlates with taurine and the bone formation marker osteocalcin during early postnatal life and aging in humans. (AF) Pearson correlation scatter plots between (A and D) serum B12 and taurine, (B and E) B12 and osteocalcin, and (C and F) osteocalcin and taurine in (AC) children of B12-deficient mothers (red symbols n = 5) and of healthy mothers (white symbols n = 7) and in (DF) aged B12-deficient subjects (red symbols n = 8) and healthy controls (white symbols n = 10). Pearson R as well as P values are shown. See also Supplemental Figure 8.

Analysis of aged patients with B12 deficiency showed significantly decreased levels of taurine and osteocalcin (Supplemental Figure 8C). Further correlation analysis using all samples from aged subjects showed significant positive correlation between variation in serum B12 and taurine (R = 0.707 P = 0.001), B12 and osteocalcin (R = 0.946, P = 2.6 × 10 –9 ), and taurine and osteocalcin (R = 0.699 P = 0.0012) levels (Figure 8, D–F). These data provide further support for a physiological role of B12 in regulating taurine synthesis and bone formation in humans. However, the human sample size in our studies was small follow-up controlled clinical trials correlating the markers of the B12/taurine axis with bone formation using larger sample sizes would be needed.

Our present findings uncovered an unanticipated regulation of growth and bone mass through a vitamin. These findings shift the focus to maternal nutritional milieu as a major player in skeletogenesis and provide conclusive evidence that maternally derived B12 regulates growth and bone accrual in the offspring via the GH/IGF1 axis.

Our studies demonstrating an effect of B12 derived from the mother on postnatal skeletogenesis of the offspring increases the repertoire of maternal effects on organ growth and bone mass. To our knowledge, B12 is the first vitamin of maternal origin whose deficiency or excess specifically manifests its consequence on postweaning bone formation. Our conclusion that alterations in maternal ability to transfer B12 to the offspring regulated growth and bone mass acquisition was based on 2 lines of evidence. First, only pups born from homozygous Gif –/– mothers, not those born from heterozygous Gif +/– mothers, had early-onset growth retardation and osteoporosis. Second, a single injection of B12 to the homozygous mothers was able to transfer sufficient B12 to their progeny and prevented the development of growth retardation and osteoporosis. The onset of growth retardation in the offspring only after weaning indicated that only minimal B12 is required during critical periods of pregnancy for functional folate/methionine metabolism. A recent study reported that mutated methionine synthase reductase (Mtrr), which activates the MTR/B12 complex, caused transgenerational embryonic abnormalities and/or lethality that were much more severe than our B12-deficient mouse model ( 38 ). On the other hand, in our present study, the higher stability of B12 overcame this neonatal lethality and allowed us to investigate the effects of deficiency in methionine/folate metabolism on mouse postnatal growth and metabolism. Together, our results indicate that modifications in methionine/folate metabolism during development or postnatal life will have long-lasting consequences on growth and bone homeostasis.

B12 deficiency results in ablation in the activity of the enzymes MTR and MUT in mammals and, consequently, accumulation of their substrates, including homocysteine and methylmalonic acid (MMA), respectively. The observation that increased homocysteine and MMA levels in serum and tissues upon B12 deficiency have diverse effects on multiple intracellular pathways led us to investigate whether the B12 deficiency–mediated growth retardation and low bone mass observed during P21–P46 was caused by their accumulation. Administration of homocysteine (50 mg/kg/d i.p.) or MMA (1 μmol/g BW/d i.p.) to WT mice beginning at P21 for 4 weeks, the period during which we observed growth retardation upon B12 deficiency, had no effect on the growth or bone mass of WT animals (Supplemental Figure 3, E and F). These results were consistent with prior reports that postnatal administration of MMA does not affect growth of rats ( 39 ). Our findings suggest that the growth retardation and low bone mass in B12-deficient animals observed from P21 was not caused by accumulation of MMA or homocysteine, but rather by downregulated production of other B12-dependent downstream metabolites. However, it is possible that the dose and duration of treatments used herein did not saturate the intracellular compartments of MMA or homocysteine, and that further increasing their levels, or levels of other metabolites, for longer durations may affect cellular processes underlying growth. Finally, our finding that taurine administration rescued the molecular and cellular abnormalities observed upon B12 deficiency suggested that taurine synthesis lies upstream of the metabolites perturbed upon B12 deficiency.

Analysis of Gif mutants for 2 generations showed that maternally derived B12 profoundly regulated GH-dependent processes in the offspring. Investigation into the GH resistance of B12-deficient mutants lead to the identification of major liver metabolic pathways underlying this resistance. Multiple lines of evidence provide credence to our assertion that the decrease in bone mass and organ growth observed upon B12 deficiency is caused, at least in part, by GH resistance. First, B12-deficient mice closely phenocopy Ghr –/– and Stat5 –/– mice ( 32 , 40 ). These mutant mice have similar changes in hepatic gene expression, including reduced Igf1, Igfals, Mup1, Slco1a1, and Hsd3b5 mRNA and increased Igfbp1. Second, both B12-deficient and Stat5 –/– mice show a major decrease in serum IGF1 levels. Third, there was a 10% reduction in serum total protein content in B12-deficient mice (40.1 ± 5.1 g/l, versus 50.4 ± 4.0 g/l in WT), an observation consistent with a major reduction in GH-mediated anabolic processes such as protein synthesis. Taken together, the striking similarities between B12-deficient and Stat5 –/– mice strongly support the notion that the effects of B12 deficiency, at least on growth and bone mass, are mediated by abrogation of STAT5-dependent signaling downstream of GH.

Investigation into the downstream mediators of GH action that underlie growth retardation and osteoporosis in B12-deficient animals led to the discovery of taurine as a metabolite regulated by B12 and underlying GH resistance in B12-deficient offspring. Taurine is a poorly understood amino acid whose function in body physiology has not been clear. Our demonstration that one means by which GH regulates growth and bone mass is through regulation of B12-dependent liver taurine production is supported by 3 lines of evidences. First, taurine production was regulated by GH in hepatocytes in a STAT5- and B12-dependent manner, because in the absence of either, GH was unable to increase taurine production. Second, taurine increased IGF1 synthesis from the liver and its action on osteoblasts by regulating IGF1R signaling. Third, feeding taurine to B12-deficient animals prevented their growth retardation and osteoporosis by normalizing their GH resistance through the activation of an alternate methionine synthesis pathway (BHMT). Our finding that B12 deficiency or taurine administration did not affect Igf1 expression in bone, but did so profoundly in the liver (Supplemental Figure 7, B–D), suggests that liver is much more sensitive to changes in B12 levels than bone. This is further supported by our in vitro results wherein B12 deficiency in osteoblasts did not affect their proliferation or function, whereas it led to hepatocyte dysfunction. We note that B12 deficiency or taurine-mediated rescue of bone mass was not associated with changes in mRNA levels of markers of osteoblast differentiation. It is likely that taurine affects osteoblast differentiation markers more profoundly at the level of mRNA translation/protein synthesis, a major function regulated by the GH/IGF1 axis. This explanation is supported by our finding that protein levels of OCN, a marker of osteoblast differentiation, were affected by changes in B12/taurine levels, whereas mRNA levels of Ocn were not (Supplemental Figure 7, F and G). Finally, our identification of taurine synthesis from the liver as an essential process, in the absence of which IGF1 could not rescue growth and bone mass downstream of B12, revealed an unanticipated role of taurine as an upstream regulator of IGF1 synthesis and action, further expanding the importance of IGF1 in body physiology ( 27 ).

In summary, our present findings (a) show an unanticipated regulation of growth and bone mass by maternally derived B12 through regulation of the GH/IGF1/taurine axis in the offspring (b) identify the cellular and metabolic abnormalities associated with B12 deficiency (c) show that taurine synthesis in the liver lies upstream of IGF1 synthesis and action and is necessary and sufficient to prevent the growth retardation and osteoporosis observed upon B12 deficiency and (d) demonstrate that taurine mediates its action on growth and bone mass through regulation of IGF1 synthesis from the liver and its action in bone. Together, these in vitro and in vivo results support the concept that B12 regulates growth and bone mass through the regulation of GH sensitivity in the offspring by regulating liver taurine production, and increase the importance of the role played by liver in regulating whole-body physiology ( 41 ).

Our identification of a novel gut/liver/bone endocrine axis (Figure 7J) as a regulator of growth and bone mass raises several questions. First, what are the molecular regulators of B12 absorption and the molecules that are coabsorbed with it in the body physiology? Second, are there other vitamins that affect bone formation through similar maternal mechanisms? Third, what are the other effects of B12 on body physiology? Fourth, how does taurine increase BHMT levels and activity, and are there other metabolites that can perform the same function? Finally, as suggested by the profound positive influence of B12 on bone formation, could regulation of B12 and/or its downstream effector taurine have the potential to increase bone formation to treat skeletal diseases? Addressing these questions will require further investigation into the regulatory mechanisms that surround B12 physiology and the GH/taurine axis, and may lead to the development of novel therapies to cure growth and bone diseases.

Gif –/– mice (Gif tm1aWtsi/tm1aWtsi ), generated by the Sanger Mouse Genetics Programme, carry a knockout-first allele in which a promoterless cassette including LacZ and neo was inserted in Gif, resulting in a loss-of-function allele. Gif –/– mice were generated as part of the mouse genetics project using C57BL/6N embryonic stem cells and were on a pure C57 background. WT littermate controls were used throughout the study.

For CN-B12 treatment to pregnant dams, virgin Gif –/– (F1) females were timed mated with Gif –/– (F1) male mice. Pregnant animals received either vehicle (saline) or 200 μg CN-B12 (s.c.) on 12.5 dpc. Where indicated, WT and Gif –/– (F1) offspring were treated orally with taurine (500 mg/kg/d) and/or the IGF1R antagonist OSI906 (50 mg/kg BW/d) from P16. Mice were culled at the indicated ages.

μCT and skeletal analyses

μCT analysis was performed using a Skyscan 1172 μCT system with standard software provided by the manufacturer (Skyscan). Histological analyses (Osteomeasure Analysis System Osteometrics) and skeletal preparations were performed as described previously ( 42 , 43 ). 6–12 animals were analyzed per group (see Supplemental Methods and figure legends).

Cell cultures and molecular assays

Primary osteoblasts or MC3T3 cells were cultured in B12-deficient medium or/and α-MEM, and cell proliferation (BrdU Cell Proliferation ELISA Roche Applied Science) and differentiation (SensoLyte pNPP Alkaline Phosphatase Assay kit AnaSpec Inc.) was performed as described previously ( 43 ). Western blot analysis was performed using standard methods using antibodies from Cell Signaling.

Human hepatoma HepG2 cells grown in α-MEM were stably transfected with STAT5b (SureSulencing shRNA Plasmid Kit Qiagen) or TCOL (ref:_00DA0IMFs._500A0Bty7J Origene) or with empty vector (mock) using Attractene (ShRNA) or Turbofect (plasmids).

Metabolomics and serum B12 analysis

Polar metabolites were extracted from mouse liver and cell samples, separated using Waters Acquity ultra performance liquid chromatography and, analyzed using triple quadrupole mass spectrometry. Serum B12 levels were analyzed using competitive immunoassay on a Siemens ADVIA Centaur Immunoassay analyzer. See Supplemental Methods for details.

Quantitative RT-PCR analysis

Total RNA was extracted using a Qiagen RNA extraction kit, and real-time PCR was performed using standard protocols.

Serum was prepared using BD Vacutainer SST, snap-frozen in liquid nitrogen, and stored at –80°C until analyzed. Serum GH was measured using Mouse Growth Hormone ELISA (Millipore Inc.), and IGF1 was measured using IGF1 Mouse/Human ELISA Kit (Abcam Inc.). Luciferase reporter assays were carried out using standard methods.

Children. All clinical evaluations and sample collections were performed by an experienced hematologist. Infants 3–24 months of age presenting with megaloblastic anemia and/or symptoms of B12 deficiency (weakness, failure to thrive, refusal to wean, vomiting, developmental delay, irritability, and tremor) were enrolled during their admission to the outpatient clinics of pediatrics and pediatric hematology of Kocaeli University Hospital. Infants with serum B12 levels <150 pmol/l were eligible for the study. Dietary history in the infants, especially consumption of breast milk and animal proteins (milk, yogurt, eggs, chicken, fish, and other meat products), as well as intake of vitamin pills during pregnancy/lactation in the mothers, was recorded.

Aged subjects. All samples used in this study were from the Kuopio Ischaemic Heart Disease Risk Factor Study (KIHD study), an ongoing, population-based cohort study to investigate the risk factors for coronary heart diseases, atherosclerosis, and other related outcomes in the Eastern Finnish population ( 44 ), and were donated by J. Kauhanen and T. Nurmi (University of Eastern Finland, Kuopio, Finland). Subjects with B12 levels <150 pmol/l were considered the B12-deficient group ( 45 ). According to this classification, 8 subjects in the B12-deficient group and 10 controls (age and gender matched) were included in this study. Clinical characteristics of the subjects are given in Supplemental Figure 8.

Results are given as mean ± SEM. Statistical analysis was performed by 2-tailed Student’s t test or χ 2 test. A P value less than 0.05 was considered significant.

Animal studies. All procedures performed on mice conformed to the ethical regulation guidelines of the Wellcome Trust Sanger Institute and to the guidelines of the UK Home Office (project license no. PPL80/2479).

Children. The ethical research committee of Kocaeli University Hospital approved the protocols (study no. 2013-1, site no. 12), and written consent was obtained from the adults accompanying the controls or patients.

Aged subjects. Clinical samples from aged subjects were from the previously described KIHD study ( 44 ).

The authors are grateful to the Sanger mouse facility for help with animal experiments Kevin McKinzee for μCT data generation and analysis Miep Helfrich for generous support and Vasudev Kantae, Bhargavi Carasala, and Jonathan Broomfield for metabolite measurements. V.K. Yadav dedicates this study to his mother, Bhagwanti Devi. P. Roman-Garcia and I. Quiros-Gonzalez are supported by ERA-EDTA postdoctoral fellowships (ERA LTF-78/2011 and 107/2012). This work was supported by the Wellcome Trust (grant no. 098051).

Conflict of interest: The authors have declared that no conflict of interest exists.

Reference information: J Clin Invest. 2014124(7):2988–3002. doi:10.1172/JCI72606.


Methodology

In the present study, published data after the year 2000 relating to dietary taurine roles in fish nutrition were analyzed and visualized by using a computational literature mining model. Literature text mining techniques have been widely used in bioinformatics and biomedical research due to the high efficiency of literature capture in any specific topic. The present study collected research data from data mining and filtering by “rentrez”, R package according to the title of the article, fish species, life stages, taurine supplementation and primary response (Winter 2017). Then, the collected data were carefully summarized and tabulated for analysis and visualization. Genetic databases including the National Center for Biotechnology (NCBI) gene database were used to collect gene frequencies of the TauT gene in different fish species (Lamurias and Couto 2019). To calculate the optimum dietary supplementation level, all the taurine data were entered separately and tabulated. Tabulated data were filtered to make graphs and figures. The data were expressed as mean ± SEM (standard error of the mean) and analyzed by one-way analysis of variance (ANOVA) using SPSS 23.0. The number of times taurine supplementation used according to fish species and taurine levels was visualized by using Tableau Desktop 2020.1. Articles were summarized according to fish species, life stages, living environment of the fish, best-recommended taurine level, with or without fishmeal, the primary response and the main protein sources in the diet. Also, the synergic effects of different nutrients with taurine were studied.

Properties and biosynthesis of taurine

The full chemical name of taurine is 2-aminomethane sulfonic acid. It is converted from l -cysteine after the process of oxidative enzymatic action in the biosynthesis processes in liver (Liu et al. 2017). In 1827, taurine was isolated initially by Leopold Gmelin and Friedrich Tiedemann (Seidel et al. 2018). It was originally found in bile acids of the ox (Bostaurus) and the name was derived from Taurus. As a sulfur-containing amino acid, taurine is highly abundant in most animal tissues, especially in marine animals. Plant and fungi contain very low concentrations (Sundararajan et al. 2014). Taurine is commonly found in muscle, brain, liver and kidney, and it helps to develop the functions of skeletal muscles, cardiovascular and central nervous systems, and the retina (Onsri and Srisawat 2016). In fish, taurine is synthesized in liver from methionine and cysteine. However, the ability of biosynthesis varies according to fish species. Also, it has been highlighted that taurine deficiency leads to certain inferior performance and physiological abnormalities (Shen et al. 2018). Taurine is generally considered as an essential amino acid for fish. It is required in primary situations when production is decreasing due to deficiencies or lack of ability to synthesize taurine in liver (El-Sayed 2013).

Taurine affects proteins because it has the main ability of directly interacting via an amine (NH3 + ) group (Bruździak et al. 2018). Taurine is involved in several metabolic pathways, such as methionine metabolism (Andersen et al. 2015), bile acid biosynthesis (Salze and Davis 2015), inner membrane transport (Luirink et al. 2005) and sulfur metabolism (Liu et al. 1994). It has many functions, such as bile acid synthesis, cell volume regulation, cytoprotection of the central nerve system and modulation of intracellular calcium (Ripps and Shen 2012). Normally, methionine-derived homocysteine is a sulfur source, and its condensation products with serine are converted into cysteine in animals. The major pathway of taurine biosynthesis includes several sequences of the oxidation process. Cysteine is converted into cysteine sulfinic acid by cysteine dioxygenase (CDO), and then hypotaurine is produced by cysteine sulfinic acid by cysteine sulfonate decarboxylase (CSD) followed by hypotaurine dehydrogenase and produce taurine (Fig. 1). CDO regulates the cysteine concentration, and CSD enzyme is the rate-limiting step in taurine biosynthesis. CDO and CSD are the key enzymes in the taurine biosynthesis process in the liver (Wang et al. 2014). Moreover, a membrane transporter of taurine has a critical role for transport and recycling of taurine. However, regulation of taurine biosynthesis differs according to the fish species because of the key enzyme activities, especially CDO and CSD. Those enzyme activities depend on the osmotic conditions, ontogenetic stages, hormone status and diet formulation. Taurine biosynthesis is higher in rainbow trout than Japanese flounder (Wang et al. 2016). Taurine is synthesized through a transsulfuration pathway by using aspartate aminotransferase by some freshwater fish species, such as rainbow trout and common carp (Guimaraes et al. 2018). However, the taurine biosynthesis pathway in fish is still poorly described in the literature (Salze and Davis 2015). The addition of taurine to zebrafish (Danio rerio) liver cells grown in taurine-free medium has little effect on transcription levels of the biosynthetic pathway genes for cysteine dioxygenase (CDO), cysteine sulfonate decarboxylase (CSAD) or cysteamine dioxygenase (ADO). In contrast, supplementation with taurine causes a 30% reduction in transcription levels of the taurine transporter, TauT. The importance of taurine to TauT gene expression in liver has been confirmed (Liu et al. 2017).

Taurine biosynthesis pathway. (Source: KEGG pathway map-00430, Liu et al. 2017). CDO cysteine dioxygenase type 1, CSD cysteine sulfonate decarboxylase, GLD glutamate decarboxylate, AED 2-aminoethanethiol dioxygenase

Low or absence of CSD activity in liver could lead to a lack or low capacity of taurine synthesis, especially in the juvenile stage of fish (Martins et al. 2018). Hepatic taurine concentration was marginally increased with the growth of rainbow trout. Furthermore, mRNA and CSD levels were dramatically increased with the growth of rainbow trout (Wang et al. 2015). Dietary sulfur amino acids, such as methionine and cysteine, stimulated taurine biosynthesis with increased hepatic CDO and liver taurine concentration, but not significantly affected the hepatic CSD activities in turbot (Psetta maxima) (Wang et al. 2014). Carnivorous fish have a lower capacity of taurine biosynthesis than herbivorous fish. Supplementation of dietary taurine increases the utilization of plant protein in carnivorous fish (Zhang et al. 2018). So, taurine improves the growth performance of several carnivorous fish, including turbot (Scophthalmus maximus) (Liu et al. 2018 Wei et al. 2018 Zhang et al. 2019), red sea bream (Pagrus major) (Takagi et al. 2010), Japanese flounder (P. olivaceus) (Kim et al. 2017) and yellowtail (Seriola quinqueradiata) (Khaoian et al. 2014 Nguyen et al. 2015). Therefore, taurine is a vital nutrient for the above-mentioned fish species especially in their rapid growth stage, where most CSD actions take place in the liver. So, all those properties are vitally important factors in fish nutrition.

Statistical analysis of research on fish taurine nutrition

According to the data set, more than 100 specific queries of the literature were tabulated. The research trend line was with R 2 = 0.46, and P value = 0.0018. A linear trend model is computed for the sum of the number of records given published years. The literature number was significantly increased by the year (P < 0.05). The maximum number was recorded in the year 2018 with 18 records, and the minimum number was recorded with one record in the year 2001, 2002, 2009 and 2010, respectively. There was a trend line of significantly increasing number of articles in the special field of taurine supplementation and metabolism because of the increase of research, funding, high demand of seafood as a protein source, limitation and the high price of fishmeal, an increasing number of concerns on taurine, and the previous research motivations. Japanese flounder (P. olivaceus) was the most studied fish species, followed by red sea bream, yellowtail and turbot. The numerous positive effects with few negative effects of dietary taurine supplementation on growth and metabolism in fish were recorded (Table 1). Further research is needed on certain fish and their different life stages to clarify the role of taurine and its nutritional value for other nutrient metabolism.

Growth performance

In most of the published studies, the positive effects of dietary taurine supplementation on the growth and feed utilization of fish were found, especially for the fish fed with plant protein-based diets. These fish species include white seabream (Diplodus sargus) (Magalhães et al. 2019), turbot (Liu et al. 2018 Sampath et al. 2020 Wei et al. 2018 Zhang et al. 2019), rock bream (Oplegnathus fasciatus) (Ferreira et al. 2014), common carp (Cyprinus carpio) (Abdel-Tawwab and Monier 2017), snapper (Lutjanus colorado) (Hernandez et al. 2018), black carp (Mylopharyngodon piceus) (Zhang et al. 2018) and channel catfish (Peterson and Li 2018). Furthermore, it was found that dietary methionine supplementation was inefficient in the plant-based diets to overcome the taurine deficiency for the growth performance of meagre (Argyrosomus regius). So, taurine supplementation is necessary for plant protein-based diets (Moura et al. 2018).

However, the nonresponse or negative effects of dietary taurine supplementation on fish were also found in some previous studies. Growth and feed utilization of barramundi (Lates calcarifer) were not significantly affected by taurine supplementation of the plant-based diets with 1.5% of the final taurine content (Poppi et al. 2018). Also, Kato et al. (2014) found no significant difference in growth, survival, feed intake and feed efficiency of red sea bream fed with or without taurine-supplemented diet. No significant effects of dietary taurine supplementation on growth performance were found in some other fish species, such as grass carp (Yang et al. 2013) and yellowtail (Khaoian et al. 2014). Furthermore, Hoseini et al. (2017) found negative effects on the growth performance of juvenile Persian sturgeon (Acipenser persicus) fed with taurine-supplemented diet compared to the controls without taurine supplementation. The similar negative results were found in Persian sturgeon (A. persicus) (Hoseini et al. 2017) and European sea bass (Dicentrarchus labrax) (Coutinho et al. 2017).

Based on the positive effects of dietary taurine supplementation, the results of most research suggested that optimal dietary taurine content was between 0.5 and 1.5%, whereas 1% was the most recorded value (Fig. 2). According to the data set, the statistically optimal content of dietary taurine for the growth and metabolism of fish was 0.91 ± 0.06% (the mean value) (Fig. 3). Among published articles, the juvenile stage was the most tested life stage of the fish. Some deviations from the statistically optimal dietary taurine content were observed because of the specific experimental conditions and different life stages of fish. So, even with the same fish species, the optimum taurine level has deviated according to the life stages, feed formula and the experimental conditions. Also, it has been suggested that optimum taurine level is a species-specific factor for fish. Kim et al. (2017) suggested that dietary taurine content was 0.9–1.3% for Japanese flounder fed with a fishmeal-based diet. Satriyo et al. (2017) suggested that a minimum level of 0.45% of taurine is required in the diet with washed fishmeal as a main protein source to normalize the physiological conditions of juvenile totoaba, namely green liver, low gallbladder-somatic index (GBSI), low plasma total cholesterol, low lipid digestibility, low erythrocyte turnover and low visceral fat content. With most of the cases utilizing more content than the optimal level, dietary taurine has no or negative effects on fish (Hu et al. 2018b Stuart et al. 2018 Zheng et al. 2016). So, the current knowledge about the optimum dietary taurine levels is highly important for aquaculture as well as for future research. In any case, taurine has shown species specific effects on fish nutrition. So, there were more positive effects as well as a few negative effects on certain fish species. Moreover, taurine is a critical nutrient for plant-based protein diets for fish when considering the growth performance.

Sum of number of records broken down by recommended/required taurine concentration (%). Circle size and the color show sum of the number of records

Radar plot of tested taurine-supplemented percentage with fish species and their life stages. Highlighted circle is the mean value of taurine% (0.91)

Anti-oxidative and immune effects

Taurine has anti-oxidative properties because of its effect on anti-oxidative enzymes and genes in the liver and intestine of fish (Coutinho et al. 2017). According to Zhang et al. (2018), anti-oxidative enzymes, including SOD and GSH-px, in juvenile black carp (M. piceus) were significantly increased by dietary taurine supplementation. The interactive effect of dietary taurine and glutamine gave significantly higher anti-oxidative capacity in Japanese flounder (Han et al. 2014). Also, increasing dietary methionine with taurine increased activities of CAT and GPX in the liver of European sea bass (Dicentrarchus labrax). Activities of the CAT, T-SOD, and the total anti-oxidative capacity (T-AOC) in rice field eel (Monopterus albus) were significantly increased with increasing dietary taurine levels (Hu et al. 2018b). The activities of SOD and the content of glutathione in juvenile black carp (M. piceus) were increased by dietary taurine supplementation in low fish meal diet (Zhang et al. 2018). The same results were found in some other species, such as European sea bass (Feidantsis et al. 2014) and common carp (C. carpio) (Abdel-Tawwab and Monier 2017).

Juvenile yellow catfish (Pelteobagrus fulvidraco) fed with all-plant-based protein diet containing 1.09% of taurine supplementation increased red blood cell, hemoglobin, total immunoglobulin, phagocytic index, respiratory burst and activities of SOD, GPX, CAT and lysozyme in blood (Li et al. 2016). However, when dietary fishmeal was replaced by soy protein concentrates with taurine supplementation, red blood cells, plasmatic hemoglobin and hematocrit in juvenile totoaba (Totoaba macdonaldi) were not significantly different from those fed control diet (López et al. 2015). Also, dietary taurine supplementation had no significant effects on immune parameters in white seabream (D. sargus) fed with both high and low fish meal diets (Magalhães et al. 2019). The same results were confirmed in Japanese flounder (P. olivaceus) (Han et al. 2014). Also, red seabream (P. major) fed low fish meal (22–36%) diets in low water temperatures (14.5 ± 1.95 °C) with 1% dietary supplementation had increased innate immunity compared with fish that received high levels of fish meal (45%). However, hematological and biochemical parameters were not affected by taurine supplementation (Gunathilaka et al. 2019).

So, taurine improved the anti-oxidative properties of fish by optimizing the anti-oxidative and immune-related parameters, both at protein and gene levels in the liver and intestine. These parameters include anti-oxidative enzymes (e.g., CAT, SOD and GPX), hemoglobin and total immunoglobulin levels.


Do Members of the Cat Family Need to Eat Meat?

The simple answer to that question in our post-Fall ( Genesis 3 ) world is “yes.” However, we know from Genesis 1:29–30 that Adam and Eve and all the animals were commanded to only eat “green plants/herbs” for food. The cat kind (likely included in the “beasts of the earth”— Genesis 1:24 ) was originally mandated to eat only plants thus, their bodies must have been able to obtain all the nutrients necessary from plants. The diet of Little Tyke (various grains) may have more closely resembled that of the original cat kind.

In a post-Fall world, meat is believed to be a necessary part of the cat diet to obtain nutrients such as taurine and cobalamin (Vitamin B12). Taurine is an organic acid that is synthesized from the amino acid cysteine. Amino acids are the building blocks of proteins, which are a major component of meat but not of plants (which are mainly carbohydrates, including indigestible cellulose). Cats are the only mammals known to lack the ability to synthesize taurine. Lack of taurine in a cat’s diet can lead to eye problems, hair loss, tooth decay, and heart problems. Taurine is so important that it is a required additive to cat food.

Vitamin B12 is found naturally in meat, milk, and eggs. Some animals (like herbivores) possess bacteria in their rumens or gut that synthesize B12. Vitamin B12 is involved in many aspects of cellular metabolism, especially DNA synthesis. Possibly all animals and humans originally had symbiotic bacteria in their gut that synthesized B12 (this relationship was altered following the Fall). It is also plausible that pre-Fall/pre-Flood plants had levels of taurine and B12 that met the requirements for the cat kind.2 Degenerative mechanisms, such as mutation following the Fall, or plants going extinct may have led to changes in the nutritional quality and availability of plants, making meat a necessary part of the cat kind diet.3

In 2006, a study was published regarding the effects of vegetarian diets on cats.4 The study found that all of the cats on the vegetarian diet had normal levels of cobalamin, and the majority had normal levels of taurine in their blood. Commercially available vegetarian cat food is supplemented with cobalamin and taurine, and both substances can also be obtained as a supplement that is added to the cat’s diet.

Little Tyke’s diet did not include supplements of these substances, even though they would have been present in the milk and eggs she consumed. However, milk and eggs were not given to her in quantities that could have provided sufficient amounts of these substances. Taurine and cobalamin levels were never evaluated in Little Tyke, but she was not found to suffer from any of the problems typically associated with deficiencies of these nutrients in cats. Obviously, there is still much to be learned concerning the metabolism and necessity of these nutrients in the cat kind diet.


Contents

Bile acid synthesis occurs in liver cells, which synthesize primary bile acids (cholic acid and chenodeoxycholic acid in humans) via cytochrome P450-mediated oxidation of cholesterol in a multi-step process. Approximately 600 mg of bile salts are synthesized daily to replace bile acids lost in the feces, although, as described below, much larger amounts are secreted, reabsorbed in the gut and recycled. The rate-limiting step in synthesis is the addition of a hydroxyl group of the 7th position of the steroid nucleus by the enzyme cholesterol 7 alpha-hydroxylase. This enzyme is down-regulated by cholic acid, up-regulated by cholesterol and is inhibited by the actions of the ileal hormone FGF15/19. [2] [3]

Prior to secreting any of the bile acids (primary or secondary, see below), liver cells conjugate them with either glycine or taurine, to form a total of 8 possible conjugated bile acids. These conjugated bile acids are often referred to as bile salts. The pKa of the unconjugated bile acids are between 5 and 6.5, [4] and the pH of the duodenum ranges between 3 and 5, so when unconjugated bile acids are in the duodenum, they are almost always protonated (HA form), which makes them relatively insoluble in water. Conjugating bile acids with amino acids lowers the pKa of the bile-acid/amino-acid conjugate to between 1 and 4. Thus conjugated bile acids are almost always in their deprotonated (A-) form in the duodenum, which makes them much more water-soluble and much more able to fulfil their physiologic function of emulsifying fats. [8] [9]

Once secreted into the lumen of the intestine, bile salts are modified by gut bacteria. They are partially dehydroxylated. Their glycine and taurine groups are removed to give the secondary bile acids, deoxycholic acid and lithocholic acid. Cholic acid is converted into deoxycholic acid and chenodeoxycholic acid into lithocholic acid. All four of these bile acids recycled, in a process known as enterohepatic circulation. [2] [3]

As amphipathic molecules with hydrophobic and hydrophilic regions, conjugated bile salts sit at the lipid/water interface and, above the right concentration, form micelles. [9] The added solubility of conjugated bile salts aids in their function by preventing passive re-absorption in the small intestine. As a result, the concentration of bile acids/salts in the small intestine is high enough to form micelles and solubilize lipids. "Critical micellar concentration" refers to both an intrinsic property of the bile acid itself and amount of bile acid necessary to function in the spontaneous and dynamic formation of micelles. [9] Bile acid-containing micelles aid lipases to digest lipids and bring them near the intestinal brush border membrane, which results in fat absorption. [6]

Synthesis of bile acids is a major route of cholesterol metabolism in most species other than humans. The body produces about 800 mg of cholesterol per day and about half of that is used for bile acid synthesis producing 400–600 mg daily. Human adults secrete between 12-18 g of bile acids into the intestine each day, mostly after meals. The bile acid pool size is between 4–6 g, which means that bile acids are recycled several times each day. About 95% of bile acids are reabsorbed by active transport in the ileum and recycled back to the liver for further secretion into the biliary system and gallbladder. This enterohepatic circulation of bile acids allows a low rate of synthesis, only about 0.3g/day, but with large amounts being secreted into the intestine. [5]

Bile acids have other functions, including eliminating cholesterol from the body, driving the flow of bile to eliminate certain catabolites (including bilirubin), emulsifying fat-soluble vitamins to enable their absorption, and aiding in motility and the reduction of the bacteria flora found in the small intestine and biliary tract. [5]

Bile acids have metabolic actions in the body resembling those of hormones, acting through two specific receptors, the farnesoid X receptor and G protein-coupled bile acid receptor/TGR5. [7] [10] They bind less specifically to some other receptors and have been reported to regulate the activity of certain enzymes [11] and ion channels [12] and the synthesis of diverse substances including endogenous fatty acid ethanolamides. [13] [14]

Bile salts constitute a large family of molecules, composed of a steroid structure with four rings, a five- or eight-carbon side-chain terminating in a carboxylic acid, and several hydroxyl groups, the number and orientation of which is different among the specific bile salts. [1] The four rings are labeled A, B, C, and D, from the farthest to the closest to the side chain with the carboxyl group. The D-ring is smaller by one carbon than the other three. The structure is commonly drawn with A at the left and D at the right. The hydroxyl groups can be in either of two configurations: either up (or out), termed beta (β often drawn by convention as a solid line), or down, termed alpha (α displayed as a dashed line). All bile acids have a 3-hydroxyl group, derived from the parent molecule, cholesterol, in which the 3-hydroxyl is beta. [1]

The initial step in the classical pathway of hepatic synthesis of bile acids is the enzymatic addition of a 7α hydroxyl group by cholesterol 7α-hydroxylase (CYP7A1) forming 7α-hydroxycholesterol. This is then metabolised to 7α-hydroxy-4-cholesten-3-one. There are multiple steps in bile acid synthesis requiring 14 enzymes in all. [3] These result in the junction between the first two steroid rings (A and B) being altered, making the molecule bent in this process, the 3-hydroxyl is converted to the α orientation. The simplest 24-carbon bile acid has two hydroxyl groups at positions 3α and 7α. This is 3α,7α-dihydroxy-5β-cholan-24-oic acid, or, as more usually known, chenodeoxycholic acid. This bile acid was first isolated from the domestic goose, from which the "cheno" portion of the name was derived (Greek: χήν = goose). The 5β in the name denotes the orientation of the junction between rings A and B of the steroid nucleus (in this case, they are bent). The term "cholan" denotes a particular steroid structure of 24 carbons, and the "24-oic acid" indicates that the carboxylic acid is found at position 24, at the end of the side-chain. Chenodeoxycholic acid is made by many species, and is the prototypic functional bile acid. [2] [3]

An alternative (acidic) pathway of bile acid synthesis is initiated by mitochondrial sterol 27-hydroxylase (CYP27A1), expressed in liver, and also in macrophages and other tissues. CYP27A1 contributes significantly to total bile acid synthesis by catalyzing sterol side chain oxidation, after which cleavage of a three-carbon unit in the peroxisomes leads to formation of a C24 bile acid. Minor pathways initiated by 25-hydroxylase in the liver and 24-hydroxylase in the brain also may contribute to bile acid synthesis. 7α-hydroxylase (CYP7B1) generates oxysterols, which may be further converted in the liver to CDCA. [2] [3]

Cholic acid, 3α,7α,12α-trihydroxy-5β-cholan-24-oic acid, the most abundant bile acid in humans and many other species, was discovered before chenodeoxycholic acid. It is a tri-hydroxy-bile acid with 3 hydroxyl groups (3α, 7α and 12α). In its synthesis in the liver, 12α hydroxylation is performed by the additional action of CYP8B1. As this had already been described, the discovery of chenodeoxcholic acid (with 2 hydroxyl groups) made this new bile acid a "deoxycholic acid" in that it had one fewer hydroxyl group than cholic acid. [2] [3]

Deoxycholic acid is formed from cholic acid by 7-dehydroxylation, resulting in 2 hydroxyl groups (3α and 12α). This process with chenodeoxycholic acid results in a bile acid with only a 3α hydroxyl group, termed lithocholic acid (litho = stone) having been identified first in a gallstone from a calf. It is poorly water-soluble and rather toxic to cells. [2] [3]

Different vertebrate families have evolved to use modifications of most positions on the steroid nucleus and side-chain of the bile acid structure. To avoid the problems associated with the production of lithocholic acid, most species add a third hydroxyl group to chenodeoxycholic acid. The subsequent removal of the 7α hydroxyl group by intestinal bacteria will then result in a less toxic but still-functional dihydroxy bile acid. Over the course of vertebrate evolution, a number of positions have been chosen for placement of the third hydroxyl group. Initially, the 16α position was favored, in particular in birds. Later, this position was superseded in a large number of species selecting the 12α position. Primates (including humans) utilize 12α for their third hydroxyl group position, producing cholic acid. In mice and other rodents, 6β hydroxylation forms muricholic acids (α or β depending on the 7 hydroxyl position). Pigs have 6α hydroxylation in hyocholic acid (3α,6α,7α-trihydroxy-5β-cholanoic acid), and other species have a hydroxyl group on position 23 of the side-chain.

Ursodeoxycholic acid was first isolated from bear bile, which has been used medicinally for centuries. Its structure resembles chenodeoxycholic acid but with the 7-hydroxyl group in the β position. [1]

Obeticholic acid, 6α-ethyl-chenodeoxycholic acid, is a semi-synthetic bile acid with greater activity as FXR agonist which is undergoing investigation as a pharmaceutical agent.

Bile acids also act as steroid hormones, secreted from the liver, absorbed from the intestine and having various direct metabolic actions in the body through the nuclear receptor Farnesoid X receptor (FXR), also known by its gene name NR1H4. [15] [16] [17] Another bile acid receptor is the cell membrane receptor known as G protein-coupled bile acid receptor 1 or TGR5. Many of their functions as signaling molecules in the liver and the intestines are by activating FXR, whereas TGR5 may be involved in metabolic, endocrine and neurological functions. [7] [18]

Regulation of synthesis Edit

As surfactants or detergents, bile acids are potentially toxic to cells, and so their concentrations are tightly regulated. Activation of FXR in the liver inhibits synthesis of bile acids, and is one mechanism of feedback control when bile acid levels are too high. Secondly, FXR activation by bile acids during absorption in the intestine increases transcription and synthesis of FGF19, which then inhibits bile acid synthesis in the liver. [19]

Metabolic functions Edit

Emerging evidence associates FXR activation with alterations in triglyceride metabolism, glucose metabolism, and liver growth. [7] [20] [18]

Other interactions Edit

Bile acids bind to some other proteins in addition to their hormone receptors (FXR and TGR5) and their transporters. Among these protein targets, the enzyme N-acyl phosphatidylethanolamine-specific phospholipase D (NAPE-PLD) generates bioactive lipid amides (e.g. the endogenous cannabinoid anandamide) that play important roles in several physiological pathways including stress and pain responses, appetite, and lifespan. NAPE-PLD orchestrates a direct cross-talk between lipid amide signals and bile acid physiology. [13]

Hyperlipidemia Edit

As bile acids are made from endogenous cholesterol, disruption of the enterohepatic circulation of bile acids will lower cholesterol. Bile acid sequestrants bind bile acids in the gut, preventing reabsorption. In so doing, more endogenous cholesterol is shunted into the production of bile acids, thereby lowering cholesterol levels. The sequestered bile acids are then excreted in the feces. [21]

Cholestasis Edit

Tests for bile acids are useful in both human and veterinary medicine, as they aid in the diagnosis of a number of conditions, including types of cholestasis such as intrahepatic cholestasis of pregnancy, portosystemic shunt, and hepatic microvascular dysplasia in dogs. [22] Structural or functional abnormalities of the biliary system result in an increase in bilirubin (jaundice) and in bile acids in the blood. Bile acids are related to the itching (pruritus) which is common in cholestatic conditions such as primary biliary cirrhosis (PBC), primary sclerosing cholangitis or intrahepatic cholestasis of pregnancy. [23] Treatment with ursodeoxycholic acid has been used for many years in these cholestatic disorders. [24] [25]

Gallstones Edit

The relationship of bile acids to cholesterol saturation in bile and cholesterol precipitation to produce gallstones has been studied extensively. Gallstones may result from increased saturation of cholesterol or bilirubin, or from bile stasis. Lower concentrations of bile acids or phospholipids in bile reduce cholesterol solubility and lead to microcrystal formation. Oral therapy with chenodeoxycholic acid and/or ursodeoxycholic acid has been used to dissolve cholesterol gallstones. [26] [27] [28] Stones may recur when treatment is stopped. Bile acid therapy may be of value to prevent stones in certain circumstances such as following bariatric surgery. [29]

Bile acid diarrhea Edit

Excess concentrations of bile acids in the colon are a cause of chronic diarrhea. It is commonly found when the ileum is abnormal or has been surgically removed, as in Crohn's disease, or cause a condition that resembles diarrhea-predominant irritable bowel syndrome (IBS-D). This condition of bile acid diarrhea/bile acid malabsorption can be diagnosed by the SeHCAT test and treated with bile acid sequestrants. [30]

Bile acids and colon cancer Edit

Bile acids may have some importance in the development of colorectal cancer. [31] Deoxycholic acid (DCA) is increased in the colonic contents of humans in response to a high fat diet. [32] In populations with a high incidence of colorectal cancer, fecal concentrations of bile acids are higher, [33] [34] and this association suggests that increased colonic exposure to bile acids could play a role in the development of cancer. In one particular comparison, the fecal DCA concentrations in Native Africans in South Africa (who eat a low fat diet) compared to African Americans (who eat a higher fat diet) was 7.30 vs. 37.51 nmol/g wet weight stool. [35] Native Africans in South Africa have a low incidence rate of colon cancer of less than 1:100,000, [36] compared to the high incidence rate for male African Americans of 72:100,000. [37]

Experimental studies also suggest mechanisms for bile acids in colon cancer. Exposure of colonic cells to high DCA concentrations increase formation of reactive oxygen species, causing oxidative stress, and also increase DNA damage. [38] Mice fed a diet with added DCA mimicking colonic DCA levels in humans on a high fat diet developed colonic neoplasia, including adenomas and adenocarcinomas (cancers), unlike mice fed a control diet producing one-tenth the level of colonic DCA who had no colonic neoplasia. [39] [40]

The effects of ursodeoxycholic acid (UDCA) in modifying the risk of colorectal cancer has been looked at in several studies, particularly in primary sclerosing cholangitis and inflammatory bowel disease, with varying results partly related to dosage. [41] [42] Genetic variation in the key bile acid synthesis enzyme, CYP7A1, influenced the effectiveness of UDCA in colorectal adenoma prevention in a large trial. [43]

Dermatology Edit

Bile acids may be used in subcutaneous injections to remove unwanted fat (see Mesotherapy). Deoxycholic acid as an injectable has received FDA approval to dissolve submental fat. [44] Phase III trials showed significant responses although many subjects had mild adverse reactions of bruising, swelling, pain, numbness, erythema, and firmness around the treated area. [45] [46]


DISCUSSION

These studies examined several biochemical and physiological mechanisms that may regulate taurine synthesis and accumulation in astrocytes. From our data, we calculate a mean rate of taurine production of 19 pmol ⋅ mg protein −1 ⋅ min −1 by dividing the increase in total culture taurine over 48 h by the average protein content per dish (0.21 mg). This value is similar to that measured from the conversion of extracellular [ 35 S]cysteine to cellular taurine. This rate of taurine synthesis from extracellular cysteine would permit complete replacement of astrocyte taurine in 2 days. In contrast, degradation is <3% over a 48-h period. Our direct measurements of [ 35 S]taurine production from [ 35 S]cysteine and quantification of CDO and CSD activities represent the first demonstration that cultured astrocytes are capable of synthesizing taurine from extracellular cysteine. These results support the glial localization of CSD shown by Almarghini et al. (1) and suggest astrocytes are major contributors to organic osmolyte synthesis in the brain. Comparing relative rates of cysteine accumulation and CDO and CSD activities suggests CDO is rate limiting for taurine synthesis. Taurine synthesis from extracellular cysteine is enhanced if turnover of the GSH pathway is eliminated, suggesting catabolism of endogenous unlabeled GSH contributes to the pool of cysteine utilized by CDO. Finally, this rate of taurine synthesis from extracellular cysteine is supported by a robust rate of cysteine accumulation.

Similar to our previous data (6), total culture taurine (medium + cells) and cellular taurine concentration increase following exposure to hyperosmotic medium. The calculated rate of increase of total culture taurine over the 48-h experimental period for cells in 450 mosmol/kg medium is 26 pmol ⋅ mg protein −1 ⋅ min −1 . This value is 37% higher than that calculated for control cells maintained in 300 mosmol/kg medium (18 pmol ⋅ mg protein −1 ⋅ min −1 ). By comparison, maximal taurine degradation in isosmotic conditions (1.3% per day) is 0.24 pmol ⋅ mg protein −1 ⋅ min −1 . Thus inhibition of taurine degradation by hyperosmotic exposure cannot explain the increase in total culture taurine. These results strongly suggest hyperosmotic exposure leads to increased taurine synthesis. A similar increase in taurine synthesis of astrocytes in situ would contribute to the elevated taurine content observed in brains of animals exposed to systemic hyperosmolality (4, 45). Although intracellular taurine concentrations increase in our cell culture system, a significant portion of the total culture taurine appears in the extracellular fluid, a volume much larger than the intracellular space. Similar efflux of newly synthesized taurine in situ would rapidly elevate the extracellular taurine concentration, thus further limiting cellular taurine loss and increasing cellular reuptake.

Taurine is a major contributor to the increase in total brain amino acids in animal models of hypernatremia (4, 44), as well as in astroglial cultures exposed to hyperosmotic medium (25, 36). This increase in situ may be due to increased synthesis, increased influx, or decreased transport of taurine out of brain parenchyma. In the present studies, total culture taurine content was increased for each hyperosmotic condition, indicating taurine synthesis is accelerated throughout the range of osmolalities tested. We anticipate astroglial taurine synthesis is similarly increased in comparable models of hypernatremia, in which experimental serum osmolalities range from 335 to 400 mosmol/kg (4, 19, 44).

Taurine flux from blood to brain does not increase during acute hyperosmotic hypernatremia (40). Enhanced taurine uptake (6, 36), and decreased taurine efflux by astrocytes in situ (6), may cause intracellular taurine sequestration however, these changes in cellular transport cannot directly increase brain taurine content. Increased glial accumulation of taurine may explain the decrease in extracellular taurine concentration described in hyperosmotic animals (18). Depending on the mechanism of brain taurine efflux, this decrease in extracellular taurine concentration may lead to decreased efflux. The contribution that changes in de novo synthesis make to the increase in total brain taurine must await quantitative data regarding the rate of taurine efflux across the blood-brain barrier of normal and hypernatremic animals.

In previous studies, we have shown that enhanced taurine uptake and inhibition of the taurine efflux pathway contribute to elevated intracellular taurine concentrations following prolonged (>24 h) hyperosmotic exposure (6). The results presented here suggest enhanced de novo taurine synthesis also occurs with hyperosmotic exposure. However, activities of key enzymes in the taurine biosynthetic pathway are not altered by hyperosmotic exposure. In addition, no change in CDO activity was measured when the reaction mixture osmolality was raised to match that which the cells experienced during the experimental treatment period, suggesting that enzyme activity is not directly altered by hyperosmotic conditions inside the cell. Although these data indicate that the quantity of enzyme is unaltered by hyperosmolality, concentrations of substrates or enzymatic cofactors or enzyme phosphorylation states (41) not measured in these studies may be altered by the hyperosmotic treatment and thus may modify taurine biosynthesis in the cell. In particular, elevated intracellular cysteine concentrations caused by cell shrinkage and increased uptake would enhance, by kinetic means, the reaction velocity of CDO.

CDO activity is highly dependent on the concentration of cysteine in the reaction mixture. Thus taurine synthesis may be regulated by the substrate availability from the extracellular space or mobilization of intracellular cysteine pools. Cellular cysteine levels increased from 4.2 to 35.6 nmol/mg protein during hyperosmotic exposure (Fig.1A), corresponding to intracellular cysteine concentrations ranging from <1 mM in isosmotic culture medium at 0 h to 5.6 mM in cultures treated in 450 mosmol/kg medium for 8 h. From Fig. 3, this elevation in cysteine concentration should increase CDO activity over sixfold. However, the production of intracellular taurine from extracellular cysteine is not increased in hyperosmotically treated cells. Conversion of extracellular [ 35 S]cysteine to [ 35 S]taurine is probably not limited by the cysteine influx rate, since, in all experimental conditions, cysteine uptake is rapid enough to equilibrate the radiolabel with the total intracellular cysteine content within the 60-min reaction period.

Cysteine utilized by CDO may be derived from extracellular or intracellular pools. We found an increased synthesis rate of taurine from extracellular cysteine when the γ-glutamyl cycle was inhibited by BSO and the remaining GSH was eliminated by CDNB, suggesting cysteine derived from GSH catabolism is shunted to taurine biosynthesis. Conversely, cysteine taken up from the extracellular space may be preferentially used by the γ-glutamyl cycle for GSH synthesis or another pathway. However, even with the GSH pathway eliminated, hyperosmotic treatment did not enhance taurine synthesis from extracellular cysteine.

Mechanisms surrounding the transient changes in cysteine and GSH contents in control (300 mosmol/kg) cultures cannot be explained by the results of this study. In previous experiments, we have shown significant decreases in intracellular taurine levels following a medium change (6). This decrease may be due to net efflux into fresh culture medium containing low concentrations of taurine or swelling-induced efflux caused by decreased osmolality of fresh growth medium. The reason for the increases in cysteine and GSH contents after 1 and 8 h is less apparent but may be a cellular response to the medium change. Nonetheless, taurine, cysteine, and GSH levels in control cells all returned to initial values by 24–48 h.

The transport system responsible for cysteine accumulation is not clear from our data, as both a sodium-independent, cystine-specific pathway and a faster sodium-dependent pathway for cysteine transport have been described in cultured astrocytes (8, 35). Although both cysteine and cystine may be present in the influx medium, Bannai (2) indicates that 90% or more of the substrate is present in the reduced form (cysteine) at the pH used during these experiments. Increased uptake in hyperosmotic conditions may be due to activation of one of these transporters, or, alternatively, increased concentrations of NaCl in the hyperosmotic experimental culture medium and influx incubation medium may enhance sodium-dependent cysteine uptake. Sodium dependence may contribute to the osmolality dependence of cysteine uptake at 1 and 8 h of hyperosmotic exposure but cannot account for decreases in accumulation that occur after 8 h. The transient increase in cysteine transport induced by hyperosmolality is similar to that described for taurine accumulation in these and other cell types (6, 28, 39). Additional experiments are necessary to describe the sodium dependence of cysteine uptake and the mechanisms responsible for increased transport rates in hyperosmotically treated astrocytes.

Elevations in cellular cysteine match the time course of increases in cysteine uptake following exposure to hyperosmotic conditions, suggesting enhanced rates of cysteine uptake directly lead to elevated intracellular cysteine levels. Because accumulation was measured with a high extracellular cysteine concentration, changes in uptake rate represent alterations in the maximal velocity of the transporter. Similar increases in uptake would also be expected at the lower physiological concentrations of cysteine that are likely to occur in situ. However, with the high extracellular concentration of cysteine used in these uptake experiments, any change in uptake due to an alteration in the transporterKm would not be detected. Sagara et al. (34, 35) and Bannai and Tateishi (3) showed that astrocyte GSH levels depend on the concentrations of cysteine or cystine in the culture medium. At all time points, astrocyte GSH contents are at least two times larger than the cellular free cysteine pool. In addition, cysteine uptake rates return to control values by 24 h while cellular cysteine levels remain elevated. This difference may represent a lag period between the initial accumulation and subsequent metabolism of cysteine to other cellular compounds.

Total γ-GT activity is not altered by exposing astrocytes to hyperosmotic culture conditions, but the enzyme rate of reaction within the cell may be decreased directly by hyperosmolality. These results may explain increases measured in GSH contents and suggest amino acid transport mediated by γ-GT turnover (23) does not contribute to the observed accumulation of amino acids during exposure to hyperosmotic conditions (25). In addition, our data indicate that the increase in cellular cysteine in hyperosmotic astrocytes is not due to enhanced metabolism of GSH. Osmolality dependence of enzymatic activity also has been described for pyruvate kinase (EC 2.7.1.40), glucose-6-phosphate dehydrogenase (EC 1.1.1.49), and LDH (EC 1.1.1.27), as well as others, and may be related to changes in protein conformation due to altered ionic strength (38, 48). In addition, the synthesis of sorbitol, another organic osmolyte, has been shown to increase in renal cells subjected to increased osmolality due to an enhancement of aldose reductase (EC 1.1.1.21) activity (43).

In summary, the accumulation and maintenance of high levels of intracellular taurine in astrocytes occur by several mechanisms. We previously reported that rapid taurine accumulation is enhanced by hyperosmotic exposure, whereas high intracellular taurine concentrations are maintained by decreases in taurine efflux (6). The present results suggest intracellular cysteine concentrations regulate taurine synthesis through the kinetic properties of CDO. During hyperosmotic exposure, increased levels of intracellular cysteine are metabolized to GSH and to taurine via CDO. Cellular GSH may be another accessible pool of cellular cysteine in astrocytes as it is in other cell types (11). This mechanism is supported, in part, by the increased rate of taurine synthesis from extracellular cysteine in GSH-depleted cells and the decline of cellular GSH during enhanced taurine synthesis in hyperosmotic cells.

We sincerely thank Dr. Martha Stipanuk and her laboratory staff for their generous assistance and advice on cysteine metabolism. We also thank the Wright State University Biomedical Science Program and the Department of Emergency Medicine for financial assistance.


The `yin and yang' of cytoprotection

Are stabilization and counteraction simply another aspect of compatibility,as it is often portrayed? Not necessarily. At its inception, the`counteracting-osmolytes' hypothesis proposed that a mixture of urea and methylamine is more beneficial than either solute alone, since a methylamine such as TMAO might `overstabilize' proteins, e.g. making them too rigid for optimal function or causing them to precipitate(Yancey et al., 1982). This concept has not received much attention, but there is evidence supporting it,as follows.

Strong stabilizers such as TMAO and trehalose appear to be high in organisms only when there is a perturbant present (e.g. urea, pressure, high temperature). The pattern of increasing TMAO with depth in marine animals(Fig. 4A) illustrates this: if high TMAO is beneficial to deep-sea animals, why isn't it used more extensively by (non-ureosmotic) shallow animals? As another example, the mammalian renal medulla appears to regulate one of its methylamine osmolytes,GPC, to maintain a constant ratio to urea, rather than for osmotic stress alone (Peterson et al., 1992). Again, if this methylamine is a simple compatible solute, why not use it at high levels under all water-stress conditions? Perhaps the costs of synthesis and retention create a tradeoff in the use of some osmolytes, but perhaps the compounds are harmful in the absence of a perturbant.

Methylamines at high concentrations can be detrimental to protein function in the absence of a perturbant, at least in vitro. For example, TMAO inhibits some enzymes (Yancey et al.,1982), and it can enhance formation of non-functional protein aggregates (Devlin et al.,2001), including β-amyloid formation(Yancey, 2001).

Using cultured renal cells, we found that adding high urea or glycine betaine alone at high concentrations to the medium greatly reduced cell growth. However, adding both partly or fully together restored normal growth(Yancey and Burg, 1990).

In yeast, high trehalose concentration, induced by temperature stress,protects enzymes at high temperatures, but rather strikingly inhibits them at normal temperatures. Yeasts that cannot eliminate their trehalose suffer when they return to normal temperatures. This has been termed `the yin and yang of trehalose' (Singer and Lindquist,1998). This phrase nicely captures the important conclusion arising from these observations, namely that some `compatible' solutes may be harmful in the absence of a perturbant.

Hypotaurine is one of the most reactive antioxidants of all known compatible solutes, and yet its use in nature at high concentrations is rare. Perhaps it is too reactive for ordinary antioxidant needs.

Some cryoprotectants such as dimethylsulfoxide and ethylene glycol, which can protect protein structure in freeze–thaw cycles, will denature proteins at higher temperatures. This may be due to the fact that hydrophobic interactions increase with temperature, such that these solutes may be excluded from proteins at low but not high temperatures(Arakawa et al., 1990).