Werner & Ingbar's The Thyroid: A Fundamental & Clinical Text, 9th Edition

7.Intracellular Pathways of Iodothyronine Metabolism

Antonio C. Bianco

P. Reed Larsen

Thyroxine (T4), the main product of the thyroid gland, is a prohormone that must be activated by deiodination to triiodothyronine (T3) in order to initiate thyroid action. This deiodination reaction occurs in the phenolic or outer ring of the T4 molecule and is catalyzed by two deiodinases, type 1 (D1) and type 2 (D2) (Fig. 7.1). In humans, approximately 80% of the T3 that is produced each day is produced by extrathyroidal deiodination of T4, demonstrating the critical importance of this pathway for thyroid hormone homeostasis. In normal humans, this occurs primarily via D2, because their serum T3 concentrations do not decline much when they are given propylthiouracil (PTU), a specific inhibitor of D1. In contrast, in rodents, both D1 and D2 contribute about equally to extrathyroidal T3 production. As a counterpoint to the activation pathway, both T4 and T3 can be irreversibly inactivated by deiodination of their tyrosyl ring (inner ring deiodination), a reaction catalyzed by either D1 or type 3 deiodinase (D3), the third member of the deiodinase group. About 40% of the T4 and nearly all of the T3 that is produced each day is deiodinated by inner ring deiodination, mostly via D3 (Fig. 7.1). Therefore, the deiodinases have the capacity to promote or terminate thyroid hormone action, constituting a critical mechanism to vary thyroid status in different organs (1).

FIGURE 7.1. Structures and interrelationships between the principal iodothyronines activated or inactivated by the selenodeiodinases.

Iodine availability is likely to have been a major factor in the evolutionary pressure leading to the selection of the deiodinases as important regulators of thyroid hormone homeostasis and biologic activity. T3 is the short-lived biologically active molecule (half-life ~1 day in humans) that is responsible for most if not all thyroid action in tissues. If T3 were the main product of thyroid secretion, its production would fluctuate according to the availability of iodine. Rather, the human thyroid secretes large amounts of T4, a long-lived molecule (half-life ~7 days) that binds to circulating proteins and accumulates in a large extrathyroidal pool, approximately 770 µg (1 µmol) in adult humans. Appropriate changes in deiodinase activity accommodate variations in iodine availability by adapting to modifications in serum T4 concentrations in order to maintain constant serum T3 concentrations (1).

STRUCTURE OF THE DEIODINASES

The three deiodinase proteins (D1, D2, and D3) are structurally similar (~50% sequence identity). All are integral membrane proteins of 29 to 33 kd, and they are especially similar in the region surrounding the active catalytic center (2,3,4). Because of inherent difficulties in the crystallization of membrane-anchored proteins, further insight into the structures of these proteins has been obtained through the use of hydrophobic cluster analysis (5), which is based on protein folding and two-dimensional transposition of sequences, allowing resolution of a sequence into its secondary structures centered on the so-defined hydrophobic clusters.

Based on this type of analysis, the three deiodinases have a single transmembrane segment, which is present near the N-terminus, and several well-conserved clusters, typical of α-helixes or β-strands, suggesting that they correspond to core secondary structures of the deiodinases. A striking common feature is the similarity of the deiodinases with various members of the thioredoxin (TRX) family, proteins that contain in their three-dimensional structure a fold defined by βαβ and ββα motifs (Fig. 7.2). The deiodinases contain extra elements, not found in the canonical thioredoxin fold, that interrupt the relationship between the βαβ and ββα motifs. These intervening sequences correspond to distinct secondary structural elements added to the canonical thioredoxin fold, a feature also present in other proteins of the thioredoxin family (6). A unique aspect of the deiodinases, however, is that one of these intervening sequences has striking similarity to α-L-iduronidase (IDUA; 47% identity with D1 and D3 and 60% with D2), a lysosomal enzyme that cleaves α-iduronic acid residues from the glycosaminoglycans heparan sulfate and dermatan sulfate (7) (Fig. 7.2). This structural similarity between the deiodinases and iduronidase may reflect the similarity of their substrates, T4 (or T3) and sulfated α-L-iduronic acid, respectively.

FIGURE 7.2. Three-dimensional structures and organization of secondary structure of the arche type thioredoxin enzyme and the rough model of the deiodinases deduced from the alignment revealed by hydrophobic cluster analysis. The transmembrane, hinge, and intervening sequences are circled. (Modified from Callebaut I, Curcio-Morelli C, et al. The iodothyronine selenodeiodinases are thioredoxin-fold family proteins containing a glycoside hydrolase-clan GH-A-like structure. J Biol Chem 2003;278:36887, with permission.)

The three-dimensional model predicts that the active center of the three deiodinases is a pocket defined by the β112 motifs of the thioredoxin fold and the IDUA-like insertion (Fig. 7.3). The most striking feature of this pocket is the presence of the rare amino acid selenocysteine, which is critical for the catalytic activity of the three deiodinases. The presence of selenocysteine was first identified when rodent D1 complementary DNA (cDNA) was found to contain UGA that encoded selenocysteine; in the vast majority of messenger RNAs (mRNAs), UGA is a stop codon (8). The cis-acting sequences that allow for the incorporation of selenocysteine rather than a stop codon consist of the seleocysteine codon (UGA) itself and a specific RNA stem-loop located in the 3′-untranslated region of the mRNA. Only in the presence of the stem-loop structure and in several trans-acting factors are UGA codons “recoded” to specify selenocysteine. The stem-loop sequence is termed the Selenocysteine (sec) insertion sequence, or SECIS, element; it is present in all three deiodinases and all other eukaryotic selenoproteins (9)

FIGURE 7.3. Schematic representation of the putative active site of deiodinases deduced from sequence alignment and from the associated modeling (see Fig. 7.2). The positions shown are those of type 2 deiodinase (D2), and the table contains the corresponding positions and residues in D1 and D3. The iduronidase (IDUA)-like insertion likely constitutes a cap that may cover the active site after ligand binding. (Modified from Callebaut I, Curcio-Morelli C, Mornon JP, et al. The iodothyronine selenodeiodinases are thioredoxin-fold family proteins containing a glycoside hydrolase-clan GH-A-like structure. J Biol Chem 2003;278:36887, with permission.)

All three deiodinases can form homodimers when transiently expressed (10,11) Fig. 7.4. This is supported by the finding of catalytic activity in the higher-molecular-weight forms (~65 kd) (12,13). However, these higher-molecular-weight forms of the deiodinases may reflect associations with other cellular proteins not primarily involved in their catalytic function, but which could, for example, regulate their half-life, intracellular transport, or subcellular localization. It is not clear that dimerization occurs or that it is necessary for function when the enzymes are expressed at endogenous levels.

FIGURE 7.4. Subcellular localization and topology of D1, D2, and D3. Left panel: Immunofluorescence confocal microscopy of HEK-293 cells transiently expressing one of the deiodinases as indicated. D1 is visible in the periphery of the cell, and does not colocalize with BiP, an endoplasmic reticulum marker visible around the nucleus. D2 is visible colocalized with BiP, whereas D3 is visible in the cell border. Right panel: Schematic representation of type 1 deiodinase (D1), D2, and D3 dimers with their respective orientations and subcellular localization. (From Toyoda N, Berry MJ, Harney JW, et al. Topological analysis of the integral membrane protein, type 1 iodothyronine deiodinase (D1). J Biol Chem 1995;270:12310; Baqui MM, Gereben B, Harney JW, et al. Distinct subcellular localization of transiently expressed types 1 and 2 iodothyronine deiodinases as determined by immunofluorescence confocal microscopy. Endocrinology 2000; 141:4309; and Baqui MM, Botero D, Gereben B, et al. Human type 3 iodothyronine selenodeiodinase is located in the plasma membrane and undergoes rapid internalization to endosomes. J Biol Chem 2003;278:1206, with per mis sion.)

TOPOLOGY AND SUBCELLULAR LOCALIZATION

The D1 and D2 monomers are integral membrane proteins oriented with a small amino-terminal extension in the extracellular space (D1) or lumen of the endoplasmic reticulum (D2), and a single transmembrane domain exiting the membrane at about position 40 (14,15). This puts the active center of both D1 and D2 in the cytosol. D3 is also an integral membrane protein, but its orientation is opposite, so that its active center is the extracellular space Fig. 7.4.

The location of mature D1 in the plasma membrane has been demonstrated in several types of cells that normally contain D1, including thyroid and kidney cells, and in cells transiently expressing the enzyme (10,16,17,18). Using confocal laser microscopy of human and mouse cells, transiently expressed FLAG-tagged D1 was also localized to the plasma membrane. D1 does not colocalize with the endoplasmic reticulum protein BiP, as does D2 (15). This plasma membrane localization has been confirmed by labeling of D1 with cell-impermeable reagents (19). D2, on the other hand, is an endoplasmic reticulum protein. Immunofluorescent confocal microscopy of human and mouse cells transiently expressing FLAG-tagged D2 revealed colocalization of D2 with BiP. Endogenously expressed D2 also colocalizes with BiP in MSTO-211H cells (20).

Using similar techniques, endogenously and transiently expressed FLAG-tagged D3 was identified in the plasma membrane. It colocalizes with the α-subunit of Na+, K+-ATPase, the early endosomal marker EEA-1, and clathrin, but not with endoplasmic-reticulum proteins. Most of the D3 is extracellular (Fig. 7.4), and its extracellular portion can be covalently labeled with a cell-impermeant probe, confirming its plasma membrane location. There is constant internalization of D3 that is blocked by sucrose-containing medium, and exposing cells to a weak base such as primaquine increases the pool of internalized D3, suggesting that D3 is recycled between plasma membrane and early endosomes. Such recycling could account for the longer half-life of D3 (12 hours), as compared with D1 (8 hours) or D2 (1 hour) (19).

A plasma membrane location for D1 could be viewed teleologically as allowing ready access of circulating T4 to the enzyme as well as facilitating the entry of the T3 produced from T4 into the extracellular fluid. The plasma membrane localization of D1 contrasts with the localization of D2 in the endoplasmic reticulum in the same cell types (15). This differential subcellular localization of D1 and D2 may explain the small contribution of T3 generated by D1 and the large contribution of T3 generated by D2 to intranuclear T3 (21,22,23,24). The extracellular location of D3 makes it readily accessible to extracellular, and therefore circulating T4 and T3, explaining its capacity for rapid inactivation of circulating T4 and T3 in patients with hemangiomas and its blockade of the access of maternal thyroid hormones to the fetus.

STRUCTURE-FUNCTION RELATIONSHIPS OF DEIODINASES AND MECHANISM OF DEIODINATION

The deiodination of iodothyronines by D1, D2, and D3 is a reductive dehalogenation that in vitro requires a reducing agent, such as dithiothreitol, as cofactor. An endogenous cofactor capable of sustaining multiple rounds of iodothyronine deiodination has not yet been identified. Based on major differences in substrate preference, reaction kinetics, and sensitivity to various inhibitors, it was assumed that the substrate-binding pockets of these enzymes were fundamentally different. However, the structural model described above predicts a conserved structure for all three deiodinases, particularly for the active center, the accuracy of which is supported by the substantial perturbations in enzyme function that result from changes of single amino acids in the binding pocket (5) (Fig. 7.3).

D1 has a relatively low affinity for T4[Michaelis-Menten constant (Km), 1,2 µM], and its catalytic activity is bisubstrate in nature, with a thiol-containing cofactor serving as the second substrate with “ping-pong” kinetics (25) Fig. 7.5.D2 and D3, on the other hand, have relatively high affinity for T4 and T3, respectively (Km, 1–4 nM), and both have sequential reaction kinetics, suggesting that the iodothyronine and the thiol-containing cofactor interact with the enzyme simultaneously before the reaction takes place (25). Regardless of the reaction kinetics, the selenocysteine residue in the active center of the three deiodinases probably acts as a nucleophile catalyzing the removal of iodine. The critical role of selenocysteine in this function was ascertained through characterization of the kinetic properties of the wild-type selenoenzymes (D1, D2, and D3) and the corresponding cysteine (Cys) mutants, which have lower affinity for the substrates and decreased turnover rates (2,11,26).

FIGURE 7.5. Mechanism of type 1 deiodinase (D1)-catalyzed thyroxine (T4) conversion to triiodothyronine (T3). The steps in the enzymatic reaction cycle at which iodoacetic acid (IAc) and propylthiouracil (PTU) are thought to inhibit catalysis are indicated. (From Leonard JL, Rosenberg IN. Thyroxine 5′-deiodinase activity of rat kidney: observations on activation by thiols and inhibition by propylthiouracil. Endocrinology 1978;103:2137, with permission.)

D1, D2, and D3 differ in their sensitivity to PTU. D1 is quite sensitive [inhibition constant (Ki), 5 µM; in vitro at 10 µM dithiothreitol], but D2 and D3 are not (Ki, >1 mM). PTU probably inhibits D1 by competing with the endogenous thiol-containing cofactor for a putative selenenyl iodide (E-Se-I) intermediate (25) Fig. 7.5. Supporting this interpretation is the fact that PTU inhibition is uncompetitive with the first D1 substrate (iodothyronine), but competitive with the second (e.g., dithiothreitol) (25). Because of the PTU insensitivity, D2- or D3-catalyzed deiodination proceeds by removal of an iodonium (I+) ion by the endogenous cofactor, resulting in an enzyme-thyronine intermediate [D2-T3 (or reverse T3)] complex and a cofactor-Se-I complex (27). In this regard, it is notable that the sequence Thr-Sec-Pro-Pro/Ser-Phe is identical in D1, D2, and D3, but that in all D1 sequences, excepting the PTU-insensitive D1 of the blue tilapia (Oreochromis aureus), the uncharged polar side chain of the Ser residue substitutes for the nonpolar side chain of Pro at position 128 (28). The existence of this natural variation prompted the creation of the Ser128Pro D1, Pro135Ser D2, and Pro146Ser D3 proteins Fig. 7.3. Remarkably, replacement of Pro135 with Ser in D2 results in a two orders of magnitude increase in Km (T4) to approximately 250 nM, approximately 10-fold lower than that of D1 for T4, and the enzyme operates with ping-pong kinetics. Furthermore, the Pro135Ser D2-catalyzed deiodination is two orders of magnitude more sensitive to PTU (Ki, 4.0 µM), although PTU inhibition is noncompetitive with dithiothreitol. Thus, the substitution of Ser for Pro135 in D2 results in changes in the enzyme that make its kinetics more similar to those of D1, indicating a critical influence of the amino acid occupying this position on enzyme function. Similar observations were made with the Pro146Ser D3 protein, which has a fivefold higher Km (T3) and is highly sensitive to inhibition by PTU (Ki, 1.0 µM). As mentioned above, Ser128Pro D1 catalyzed deiodination is resistant to PTU (Ki, >1 mM), suggesting that there is no longer an accessible E-Se-I intermediate, again illustrating the pivotal role of this position in the active center of the deiodinase molecule.

The presence of Pro in the 128/135/146 positions of D1, D2, and D3 may result in tighter binding of the substrate in the D2 (and D3) binding pockets, perhaps explaining the approximately 1000-fold higher affinity of D2 and D3 for T4, as compared with D1. This could reflect an interaction between the phenolic hydroxyl group of T4 and the Pro residue at these positions, and explain why sulfate conjugation of the phenolic hydroxyl group dramatically increases the maximum velocity (Vmax)/Km and changes the T4 deiodination site from an outer to an inner ring iodine (29).

Three conserved amino acids with charged polar side chains mark the transition between the β2-strand and the α-helix in the IDUA-like insertion, Glu/Asp155, Glu156, and His158 (D1 residues). When the invariant Glu156 (D1), Glu163 (D2), and Glu174 (D3) were replaced with Ala, the resulting enzymes had no deiodinase activity. Replacement of Glu156 with Asp in D1 supported deiodination, but with an approximately 4.5-fold higher Km (rT3), whereas a similar substitutio at position 163 in D2 did not alter the Km (T4). Thus, the acidic amino acids in this region of the deiodinase pocket are important for substrate binding or enzyme function, although the length of the side chain can vary. This is further supported by mutational studies of His at position 158 in D1 (30). Its mutation to Asn, Gln, or Phe resulted in complete loss of deiodinase activity. Replacement of the corresponding His165 in D2 with Asn also resulted in loss of deiodinase activity. According to the model, residues in this acidic pocket could interact with either the amino or the carboxyl group in the alanine side chain of the iodothyronines, a hypothesis supported by previous studies indicating that the positively charged T4 analogue (3,5,3′,5′-tetraiodothyroethylamine), which lacks a carboxyl group, is not a substrate for D1 (31). In addition, the compounds with the highest affinity for D1 (lowest apparent Km values) are those that lack positively charged functional groups (NH3+), such as tetra iodothy roacetic acid. Furthermore, the Km (T4) values for D- and L-T4 are similar (31). These results argue that the carboxyl group in the iodothyronines interacts with the NH3 group of His in the 158 position of D1 and that the other acidic residues in this pocket act to reduce the ionization of the His residue. The critical role played by the IDUA-like insertion is further strengthened by the complete loss of deiodinase activity when the conserved Trp163 in D1 and the corresponding Trp170 in D2 are replaced with Ala.

SPECIFIC PROPERTIES OF THE DEIODINASES

Type 1 Deiodinase

PTU-inhibitable, D1-catalyzed conversion of T4 to T3 supplies approximately 20% to 30% of the T3 in the serum in normal humans, but 50% or more in patients with thyrotoxicosis. D1 is the only selenodeiodinase that can function as either an outer (5′)- or inner (5)-ring iodothyronine deiodinase, D2 and D3 being exclusively outer and inner ring deiodinases, respectively (32). The molecular basis for these differences is not known.

The gene (Dio1) for human D1 is on chromosome 1 p32-p33, in a region syntenic with mouse chromosome 4, the location of mouse Dio1 (33). The complete cDNA sequences have been determined for rat, human, mouse, dog, chicken, and tilapia D1 (8,28,34,35,36,37). The size of the mRNAs for these D1s is about 2 to 2.1 kb, and all contain a UGA codon in the region encoding the active center, which is highly conserved among species. By Northern analysis, D1 is expressed in many tissues of most vertebrates, but not in amphibia (38,39,40). In rats, these include the liver, kidneys, central nervous system, anterior pituitary gland, thyroid gland, intestine, and placenta. In humans, D1 activity is notably absent from the central nervous system, but is present in the liver, kidneys, thyroid and pituitary (41,42).

Regulation of D1 Synthesis

Changes in D1 activity have been investigated in developing rats; in general, D1 activity is low in all tissues of fetal rats. It appears soon after birth in the intestine, liver, kidneys, cerebrum, cerebellum, and gonads (43). The age- related changes in D1 mRNA content are similar, indicating that the changes in activity arise at a pretranslational level. The mechanism for the age-related changes in D1 expression is unknown. The physiologic benefit of the low D1 activity in the fetus is presumably low serum T3 concentrations, thus permitting changes in intracellular T3 to be determined by the developmentally programmed changes in D2 and D3 activity (44).

Thyroid Hormone

Thyroid hormone–induced increases in D1 activity and mRNA content are well documented in rats, mice, and humans (8,45). The increases are due to increased transcription, which in the human Dio1 gene can be attributed to the presence of two thyroid hormone-response elements (TREs) in the 5′-flanking region of the gene (46,47,48). Studies in thyroid receptor (TR) knockout mice indicate that the T3-induced D1 stimulation is largely mediated by the β subtype of the receptor (49). Given this finding, the responsiveness of the human Dio1 gene to T3 would be expected to be greatest in patients with thyrotoxicosis. In fact, the D1 mRNA content of peripheral blood mononuclear cells is increased in proportion to the degree of thyrotoxicosis (42). This increase can explain the acute decrease in serum T3 concentrations that occurs in response to PTU in patients with thyrotoxicosis (50).

Cytokines

Interleukin-1 (IL-1), IL-6, tumor necrosis factor-α (TNF-α), and other cytokines are potential mediators of the alterations in thyroid function that occur in patients with severe nonthyroidal illness (see section on nonthyroidal illness in Chapter 11) (51,52,53). TNF-α, IL-1β, and interferon-γ decrease D1 activity and mRNA in rodent thyroid (FRTL5) cells (54). The effects of TNF-α have been examined in hepatocytes and HepG2 cells with contradictory results. TNF-α decreased the stimulatory effect of T3 on D1 mRNA production in HEPG2 cells, an action that is blocked by nuclear factor kappa B (55). In dispersed rat hepatocytes, IL-1βand IL-6 blocked T3 stimulation of D1 mRNA and activity; however, TNF-α had no effect (56). The effect of IL-1βwas blocked by coexpression of the nuclear steroid receptor coactivator-1 (SRC-1), but not by CREB binding protein (CBP) or CBP-associated factor (pCAF). Because IL-1 did not alter the amount of SRC-1 in hepatocytes, the effect was attributed to competition between IL-1 and T3-stimulated transcriptional events for limiting quantities of SRC-1. More recent studies describing decreases in D1 activity in liver and D2 activity in skeletal muscle and increases in D3 activity in liver and skeletal muscle in patients dying of multiple organ failure and related conditions are discussed in the section on nonthyroidal illness in Chapter 11.

Nutritional Influences on D1 Expression

A decrease in serum T3 concentrations relative to those of T4 and an increase in serum reverse T3 concentrations during fasting in humans was one of the earliest indications that the peripheral metabolism of thyroid hormones in humans was modulated by physiologic or pathophysiologic events (57). Similar changes occur in virtually all acutely ill and many chronically ill patients (58,59). Because thyroidal secretion accounts for only about 20% of daily T3production in humans, the illness-associated decrease in serum T3 concentrations must be caused, largely if not completely, by decreased T4 to T3 conversion by D1 or D2 or by increased T3 clearance by D3 (60,61).

Early studies of liver D1 activity in rats suggested that the decrease in T4 conversion to T3 that occurred during fasting might be caused by a decrease in the thiol cofactor that serves as the cosubstrate for D1-catalyzed T4 to T3 conversion (62,63). However, this cofactor has not been identified. Although rats have been studied extensively as a model for the effects of fasting (and illness) on T4 to T3 conversion in humans, they are a poor model for the effects of fasting in humans because of their low body fat content and the fact that, unlike humans, their serum TSH and T4 concentrations decrease rapidly when they are starved (64). Also, despite reduced hepatic D1 activity, total body conversion of T4 to T3 is not reduced during starvation in rats (65,66). The marked fasting- induced reduction in serum TSH, T4, and T3 concentrations (i.e., central hypothyroidism) in rats is probably due, at least in part, to leptin deficiency (67). Prefeeding rats with a high-fat diet to induce obesity results in less urinary nitrogen loss and a lesser decline in serum T4 and T3 concentrations during starvation, and serum T3 concentrations actually increase if the period of starvation is prolonged (64). In contrast, in humans, serum T3 concentrations decrease rapidly to about 50% of baseline during fasting, and they remain low for up to 3 weeks of fasting, but serum T4 and TSH concentrations change little (68).

Selenium Availability

A decrease in hepatic D1 activity of Se-deficient rats and the demonstration that D1 could be labeled with 75Se were the first clues that this trace element was critical to the function of D1 (69,70,71,72). However, the effects of Se deficiency on the synthesis of intracellular selenoproteins, such as the selenodeiodinases, depend on the tissue being examined. For example, in Se-deficient rats, thyroidal D1 activity is preserved, while that in the liver declines precipitously (73), serum T4 concentrations increase, and serum T3 concentrations do not change (74). Se deficiency also decreases D1 activity in the kidneys; this is accompanied by a decrease in D1 mRNA, which does not occur in the liver (75). Se deficiency can occur in patients receiving diets that are restricted in protein content, such as those given for phenylketonuria, and has also been found in elderly patients (76,77,78,79). In Se-deficient humans, serum T4concentrations and the serum ratio of T4 to T3 are slightly increased, but serum TSH concentrations are normal. In one endemic goiter region in Africa, there is an accompanying Se deficiency (80,81). When Se was supplied to these iodine-deficient people, their thyroid function deteriorated, as evidenced by an increase in serum TSH concentrations and a decrease in serum T3 concentrations, suggesting that the reduction in D1 activity during Se deficiency can protect against iodine deficiency, presumably by reducing inner ring deiodination of T4 or T3 (82,83).

Type 2 Deiodinase

Type 2 deiodinase is an obligate outer ring selenodeiodinase that catalyzes the conversion of T4 to T3. The Km of D2 for T4 is in the nanomolar range under in vitro conditions in the presence of 20 mM dithiothreitol. The presence of considerable D2 activity in human skeletal muscle, unexpected because it is absent in rats and mice, provides a plausible source for a substantial fraction of the extrathyroidally generated T3 in human serum (26). T4 causes a posttranslational decrease in D2 activity, due to stimulation of proteolysis of D2 via the ubiquitin-proteasome pathway (84,85,86).

Gene Structure and Chromosomal Localization

The Dio2 gene is present as a single copy located on the long arm of chromosome 14 (14q24.3) in humans (87,88). It is about 15 kb in size, and the coding region is divided into two exons by an intron of approximately 7.4 kb. The exon/intron junction is located in codon 75, and is at the same position in the human and mouse Dio2 genes (87,89,90). For the human gene, there are three transcriptional start sites, 707, 31, and 24 nucleotides 5′ to the initiator codon ATG. The longest 5′-untranslated region of human D2 mRNA contains an intron of approximately 300 bp that can be alternatively spliced (90). Other splicing variants involving the coding region have also been identified (91). The human, mouse, and rat Dio2 5′-flanking regions have been isolated and functionally characterized. All contain a functional cyclic adenosine monophosphate (cyclic AMP) response element, but only human Dio2 has binding sites for thyroid transcription factor-1 (TTF-1) (90,92,93).

D2 mRNA and Protein

Human, mouse, and chicken D2 cDNAs containing intact 3′-untranslated regions (5- to 7.5-kb) have been identified using GenBank searches and library screening. These D2 cDNAs encode functional D2 proteins, as determined by expression in Xenopus laevis oocytes (89,94,95). Rat and human D2 mRNAs are approximately 7.5 kb, and chicken cDNA is approximately 6 kb (93,94,95,96). A detailed analysis involving nuclease mapping, primer extension, and Northern blots indicated that human D2 mRNA exists as four different transcripts in thyroid, brain, and possibly other tissues (90). The longest transcript is approximately 7.5 kb, starts 708 nucleotides up-stream from the initiator ATG codon, and is the only transcript found in placenta. A shorter (approximately 7.2-kb)-minor D2 species uses the same transcriptional start site, but the approximately 300-bp intron is spliced out. Two shorter transcripts of approximately 6.8 kb, differing by only seven nucleotides, use 3′ transcriptional start sites close to the translation initiation site. It is not known whether the rat and mouse genes use the same two ma jor transcriptional start sites, but this is likely to be so for D2 in rat brain (26,90,96). The deduced amino acid sequences of the chicken, mouse, rat, and human D2 enzymes contain two selenocysteine residues. The first is in the active center of the enzyme, whereas the second is located close to the carboxy-terminus. In fish and frog D2, there is only one selenocysteine codon, which is located in the active center of the enzyme (26,89,95,96,97,98). Truncating the C-terminal amino acids, including the C-terminal selenocysteine, in human D2 has no effect on D2 enzyme kinetics or activity (99).

Tissue Distribution

In rats, D2 activity is predominantly expressed in the pituitary, brain, and brown adipose tissue (24,100,101,102,103,104. D2 activity is also present in the gonads, pineal, thymus, and uterus of rats, mammary gland of mice, and vascular smooth muscle cells in humans (43,105,106,107,108,109). High levels of D2 mRNA and activity are found in the mouse cochlea at the eighth postnatal day, suggesting a role for D2 in generating T3 for cochlear development (110). In the cerebral cortex of neonatal rats, D2 mRNA is present in astrocytes (111) and tanycytes; the latter are specialized ependymal cells lining the third ventricle that have multiple cellular processes that express D2 mRNA and that extend to the median eminence (111,112,113,114). A monosynaptic pathway has also been identified between the arcuate nucleus, which contains D2, and the paraventricular nucleus, which contains thyrotropin-releasing hormone (TRH) (115).

In humans, D2 mRNA or activity is expressed not only in vascular smooth muscle cells, but also in the thyroid, heart, brain, spinal cord, skeletal muscle, and placenta, and small amounts of D2 mRNA have been detected in the kidneys and pancreas (26,90,96,116,117,118). Thyroid tissue contains relatively more D2 mRNA than D2 activity, with the exception of thyroid tissue in patients with thyrotoxicosis caused by Graves' disease and follicular adenomas, in which both D2 mRNA and activity are present in large amounts (116). The discrepancy between D2 mRNA and activity is probably due to substrate-induced D2 ubiquitination. D2 mRNA sequences are also present in libraries from prostate, breast, and uterus, but none of these tissues have D2 activity (94). D2 mRNA or activity is present in human pituitary glands and brain tumors (116,119,120), and D2 activity has been found in human keratinocytes (121) and mesothelioma cells (20).

Regulation of D2 Synthesis

The Dio2 gene is regulated in part by a cyclic AMP-mediated pathway. Cold exposure increases D2 mRNA and activity in brown adipose tissue in rodents, and α1- or β-adrenergic antagonist agents block this effect (104,122). In isolated brown adipocytes, the increase of D2 activity during catecholamine treatment is actinomycin D sensitive (123,124,125). In addition, D2 activity in brown adipose tissue is induced by norepinephrine, isoproterenol, insulin, and glucagon, and it is inhibited by growth hormone (126,127). Cyclic AMP increases D2 activity in mesothelioma cells (20) and in rat astroglial cells (101,128), as does both nicotine and cyclic guanosine monophosphate (cyclic GMP) (129,130). As noted above, D2 mRNA and activity are increased in thyroid tissue from patients with Graves' thyrotoxicosis, and forskolin increases D2 mRNA in dispersed human thyroid cells (90,116). It is therefore not surprising that human, rat, and mouse Dio2 contains a cyclic AMP response element approximately 90 nucleotides upstream of the transcriptional start site (90,92,93,131). The promoter activity of human Dio2 increases 10-fold when cells are cotransfected with Dio2 and the α-catalytic subunit of protein kinase A. Mutation of the latter element abolishes the effect and decreases basal expression of Dio2 by approximately 90% (90).

Although there is a high level of D2 mRNA in human thyroid tissue, no D2 mRNA or activity is present in FRTL-5 rat thyroid cells, and in adult rat thyroid tissue D2 mRNA levels are very low and D2 activity is undetectable (93,116,132). Expression of the Dio2 gene in human thyroid tissue is under the control of TTF-1 but is not affected by Pax-8 (93). The human Dio2 gene has two TTF-1 binding sites, which are not present in the rat Dio2 gene, despite an overall 73% cross-species homology. The lack of these sites may explain the very low expression of D2 mRNA and activity in rat thyroid tissue.

Regulation of Degradation of D2

D2 is the critical T3-generating deiodinase due to its substantial responsiveness to physiologic signals. For example, D2 responsiveness to cyclic AMP constitutes the basis for the adrenergic stimulation of D2 activity in brown adipose tissue, and human skeletal muscle and thyroid tissue. This links D2 expression with the sympathetic nervous system and widens the spectrum of environmental and endogenous stimuli that can potentially influence adaptive T3production (see reference 1 for review).

Several transcriptional and posttranslational mechanisms have evolved to ensure tight control of tissue levels of D2, which is inherent to its homeostatic function. The D2 mRNA in higher vertebrates is more than 6 kb in length, containing long 5′ and 3′ untranslated regions. The D2 5′ untranslated regions are greater than 600 nucleotides in length, and they contain three to five short open reading frames, which reduce D2 expression by as much as fivefold (133). Alternative splicing is another mechanism that regulates the level of D2 synthesis, because mRNA transcripts similar in size to the major 6- to 7-kb D2 mRNAs, but not encoding an active enzyme, are present in both human and chicken tissues (133).

The ratios of D2 activity to D2 mRNA level in tissues vary, indicating substantial posttranslational regulation of D2 expression (134). In fact, the decisive property of D2 that characterizes its homeostatic behavior is a half-life of approximately 40 minutes that can be further reduced to approximately 25 minutes by exposure to physiologic concentrations of its substrate, T4, or extended to approximately 300 minutes when cells are grown in medium lacking T4(135,136,137,138,139,140,141,142). This constitutes a rapid, potent regulatory feedback loop that efficiently controls T3 production and intracellular T3concentrations based on how much T4 is available. The potency of the T4 in inducing loss of D2 activity mirrors the enzyme's affinity for the substrate, indicating that enzyme–substrate interaction must occur in order to induce loss of D2 activity.

At the molecular level, D2 activity is regulated by conjugation to ubiquitin, a protein of approximately 8 kd. The ubiquitinated D2 is subsequently recognized and degraded by proteasomes (143,144) (Fig. 7.6). The first evidence for this process was obtained in GH4C1 cells, in which the half-life of endogenous D2 was noted to be stabilized by MG132, a proteasome inhibitor (84). Substrate-induced loss of D2 activity was also inhibited by MG132 in these cells, indicating that both pathways affecting loss of D2 activity were mediated by the proteasomes. This implies that the loss of D2 activity is at least partially due to proteolysis of D2, a premise that was confirmed when the levels of immunoprecipitable D2 were found to parallel D2 activity, both under basal conditions and after exposure to T4 (85). In subsequent studies it became clear that D2 is ubiquitinated (86), and the various enzymes involved in this process were identified. In studies in which human D2 was expressed in yeast, Ubc6p and Ubc7p were identified as the ubiquitin conjugases involved in ubiquitination of D2 (145), and it is now clear that these conjugases play a role in ubiquitination of human D2 (146,147).

FIGURE 7.6. Representation of ubiquitination and proteasomal degradation of type 2 deiodinase (D2). Ub, ubiquitin; E1, enzyme that activates Ub; E2, Ub conjugase; Ubc6 and Ubc7, E2s involved in D2 ubiquitination; Cue1, endoplasmic reticulum–docking protein for Ubc7; asterisk in the D2 molecule, selenocysteine-containing active center; isopeptidase catalyzes D2 deubiquitination.

Fusion of the 8–amino acid FLAG sequence to the carboxyl-, but not the amino-, terminus of D2 prolongs its activity and increases the size of the ubiquitin-D2 pool by 20- to 30-fold (86), suggesting that D2 ubiquitination is reversible, because not all Ub-D2 undergoes proteolysis. Enzymatic deubiquitination of ubiquitin-D2 occurs in vitro (148) and could explain recycling in vivo. D2 was recently identified as a substrate for the deubiquitinating enzymes VDU1 and VDU2 (149). Confocal studies indicate that both VDUs colocalize with D2, itself an integral endoplasmic-reticulum membrane protein. The physical colocalization of VDU with D2 provides the opportunity for deubiquitination of D2.

VDU1-catalyzed D2 deubiquitination is an important part of the adaptive mechanism that regulates thyroid hormone action. In stimulated brown fat tissue, D2 increases intracellular T3 production, resulting in isolated tissue thyrotoxicosis (150,151,152). This is an important mechanism for cold acclimatization in rodents; mice with targeted inactivation of the D2 gene develop hypothermia and marked weight loss during cold exposure due to impaired thermogenesis in brown adipose tissue (151,153). Increased VDU1-catalyzed deubiquitination of ubiquitin-D2, and therefore rescue of D2 from proteasomal degradation, is an integral part of this mechanism. In brown adipose tissue, VDU1 mRNA levels are markedly up-regulated by cold exposure or norepinephrine, which amplifies the transcriptional increase in D2 activity, and hence T3 production increases by approximately 2.5-fold. Although ubiquitination is known to play a physiologic role in several cellular processes (154,155,156,157,158), enzyme reactivation due to deubiquitination is unusual.

The availability of a reversible ubiquitination-dependent mechanism to control the activity of D2 constitutes a biochemical and physiologic advantage that allows for rapid control of thyroid hormone activation. The finding that VDU1 and VDU2 are coexpressed with D2 in many human tissues, including brain, heart, and skeletal muscle (1,159), suggests that the importance of this mechanism may extend well beyond thermal homeostasis to include brain development, cardiac performance, glucose utilization, and energy expenditure.

Type 3 Deiodinase

D3, acting by inner ring deiodination, is the major T3- and T4-inactivating enzyme, although D1 also has some activity as an inner ring deiodinase (160). D3, which has almost exclusively inner ring deiodination activity, catalyzes the conversion of T4 to reverse T3 and the conversion of T3 to 3,3′-diiodothyronine (T2), both of which are biologically inactive Fig. 7.1. That reverse T3 and T2 do not support thyroid hormone–dependent gene expression is illustrated by the severe consumptive hypothyroidism that occurs in patients with hemangiomas, in whom tumor overexpression of D3 results in very high serum reverse T3 concentrations, and the blockade of metamorphosis that occurs in tadpoles overexpressing D3 (161,162).

D3 contributes to thyroid hormone homeostasis by protecting tissues from an excess of thyroid hormone. It was identified in monkey hepatocarcinoma cells (NCLP6E), and the first physiologic studies were performed in the central nervous systems of rats (163,164,165,166). In humans, D3 is present in not only the central nervous system, but also skin and placenta; it is also present in fetal liver and in the uterus of pregnant rats (167). The highest activity found to date is in hemangioma-type tumors in humans (162). In amphibians, D3 plays a critical role in development (168); it is present in tadpoles from premetamorphosis to the onset of the metamorphic climax, after which it declines to barely detectable levels. In mammals, D3 is critical for thyroid hormone homeostasis, because it protects the fetus from premature exposure to excessive amounts of thyroid hormone, which can result in malformations, altered growth, mental retardation, and even death. In fetal and neonatal animals, D3 expression is highly regulated in tissue-specific patterns that are likely to be critical to the coordinated regulation of thyroid hormone effects on development.

Gene Structure and Chromosomal Localization

The Dio3 gene is located on human chromosome 14q32 and mouse chromosome 12F1 (169). A unique feature of the human and mouse Dio3 gene is that it has no introns (3), which is a rarity among eukaryotes (169,170). The gene is preferentially expressed (imprinted) from the paternal allele in mice (171). The Dio3gene likely belongs to the same cluster of imprinted genes in mouse chromosome 12 and human chromosome 14, and as such it might play a role in the phenotypic abnormalities associated with uniparental disomy of those chromosomes, a condition in which gene expression is altered due to abnormal genomic imprinting (172).

Human D3 mRNA contains 2066 nucleotides. There are 220 bp in the 5′-untranslated region, an 834-bp open reading frame, and a 3′-untranslated region of 1012 bp (173). All D3 cDNAs identified to date include a selenocysteine-encoding TGA codon, as well as SECIS element in the 3′-untranslated region. There is a high degree of identity between the Dio3 gene in human and other species, particularly in the putative active center where the selenocysteine is located. The conservation of this enzyme from tadpoles to humans implies that its role in regulating thyroid hormone inactivation during embryologic development is essential. The most common form of D3 mRNA in most tissues is 2.3 kb, but there are at least four differently sized mRNAs in the central nervous system of rats, and thyroid hormone causes increases in the relative intensity of these mRNAs (174).

Tissue Distribution

D3 has been identified in various tissues in several animal species, among which rats have been studied most extensively. In adult rats, D3 is found predominantly in the central nervous system and skin. In neonatal rats, it is found not only in these tissues, but also in skeletal muscle, liver, and intestine (43,164,175,176,177,178). In particular, using in situ hybridization analysis, D3 mRNA has been identified throughout the brain in adult rats, especially in hippocampal pyramidal neurons, granule cells of the dentate nucleus, and layers II to VI of the cerebral cortex (174). It is noteworthy that these regions also contain the highest con centrations of thyroid receptors in the brain, and they have critical roles in learning, memory, and higher cognitive functions (179,180,181). Furthermore, the distribution of D3 mRNA in the central nervous system changes during the early stages of development. At postnatal day 0, D3 is selectively expressed in the bed nucleus of the stria terminalis, the preoptic area, and other areas related anatomically and functionally to the bed nucleus, such as the central amygdala; all of these areas are involved in the sexual differentiation of the brain (182). D3 expression in these areas was transient and was no longer detected at postnatal day 10. The overall pattern of distribution of D3 in the brain of rats strongly suggests that it is primarily expressed in neurons, but it is also present in astrocytes (183,184,185).

Very high levels of D3 activity and mRNA have been identified in hemangiomas in infants. In infants with very large tumors, the result is hypothyroidism, caused by very rapid deiodination of T4 and T3 (162). This syndrome, termed consumptive hypothyroidism, has also been described in an adult with a large hepatic hemangiopericytoma (186).

D3 activity has also been detected in the retina in rat fetuses and, in lesser amounts, in adults (187). In Xenopus laevis, the localized expression of D3 in the cells of the marginal zone of the retina accounts for the asymmetric growth of the retina (188). As noted above, D3 is highly expressed in the skin of adult rats (43,189), and in them skin contains more reverse T3 than any other tissue, suggesting that the high levels of D3 activity in homogenates of skin accurately reflect the activity of this enzyme in vivo (189). In this regard, T4 applied to normal human skin is largely converted to reverse T3 (190).

Large amounts of D3 are present in the placenta of rats, guinea pigs, and humans, and it is by far the predominant deiodinase present in this tissue (176,191,192,193,194). High levels of D3 are also present in the uterus of pregnant rats, initially in decidual cells and later in the single-cell layer of the epithelium (195). The levels of D3 are highest at the implantation site, nearly double the highest values obtained for placental tissue (196). In humans, the highest levels of D3 are in the endometrium (Fig. 7.7), and it is also found in fetal skin, tracheal and bronchiolar epithelium, mesothelium, and intestinal epithelium (197).

FIGURE 7.7. Type 3 deiodinase (D3) activity in the uterus and its myometrial and endometrial components. The endometrial (Endo) and myometrial (Myo) layers of the uterus from two nonpregnant women were dissected and assayed for D3 activity in the presence of 1 mM propylthiouracil. The values shown are maximum velocity, as determined by Lineweaver-Burke analysis.

Regulation of D3 Synthesis

Thyroid Hormone

Parallels between D3 activity and thyroid status have been demonstrated in several species, although the underlying molecular mechanisms remain obscure. In Xenopus laevis tadpoles, administration of T3 before the climax of metamorphosis results in a rapid and marked increase in D3 activity (39). In rats, D3 activity in the central nervous system is increased by thyroid hormone administration and decreased by hypothyroidism (164). In in situ hybridization histochemical studies, D3 gene expression within the central nervous system increased 4- to 50-fold in rats made thyrotoxic, with the greatest increase in the cerebellum. Conversely, D3 mRNA is not detectable in Northern blots of brain tissue of hypothyroid rats (174). Whether the dramatic increase in D3 mRNA in rats given T3 is due to increases in gene transcription, mRNA stabilization, or a combination of these factors is not known. In X. laevis, this T3 effect is not blocked by inhibition of protein synthesis. The promoter regions of the Dio3 gene are stimulated by T3, but the magnitude of this stimulation is modest as compared with the effect of T3 on D3 activity. Regulation of D3 activity by thyroid hormone has also been demonstrated in cultured astroglial cells. In these cells, the addition of 10 nM T3 (or T4) to the culture medium caused a slow increase in D3 activity, which reached a plateau in 48 hours (198).

Sex Steroids

D3 is expressed in the uterus, and the content increases during pregnancy. In rats, uterine D3 activity increases immediately after implantation, or artificial decidualization of the uterus in pseudopregnant rats, whereas the increases in activity are minimal in the nondecidualized uterine horn in the latter rats. In spontaneously cycling female rats, D3 activity was three to eight times higher during estrus, as compared with diestrus. Furthermore, the uterine levels of D3 activity were synergistically increased in ovariectomized rats given estradiol and progesterone in various combinations. Thus, estradiol and progesterone regulate thyroid hormone metabolism in the uterus, and the implantation process is a potent stimulus for the induction of D3 activity in this organ.

Nonthyroidal Illness

Most critically ill patients have low serum T3 and high serum reverse T3 concentrations (see section on nonthyroidal illness in Chapter 11). In patients with multiple organ system failure and other serious illnesses who died, hepatic D1 activity was low, and hepatic and skeletal muscle D3 activity was high (61). These findings suggest that T4 and T3 metabolism is altered in tissue-specific ways in illness, particularly with respect to reducing T3 formation or increasing T3 degradation. Similarly, D3 activity was increased in cardiac muscle of rats with cardiac hypertrophy and failure (199); among these rats, right ventricular D3 activity was significantly higher in those animals in which hypertrophy progressed to heart failure, as compared with the animals in which it did not. The induction of D3 in cardiac muscle would be expected to result in reduced intracellular concentrations of T3, which might reduce cardiac work and therefore help maintain cardiac compensation.

Extracellular Receptor Kinase–Activated Pathways

In cultured rat astroglial cells, factors that alter cellular processes through signaling cascades originating at the plasma membrane increase D3 activity. For example, D3 activity increases markedly and rapidly after exposure of the cells to 12-O -tetradecanoyl phorbol-13-acetate (TPA), fibroblast growth factor, epidermal growth factor, platelet-derived growth factor, and cyclic AMP analogues (200). The stimulatory effects of TPA and fibroblast growth factor on D3 mRNA and activity appear to be mediated at least partially by activation of the MEK/ERK signaling cascade (201).

NONDEIODINATIVE METABOLISM OF IODOTHYRONINES

Iodothyronines are mostly metabolized by deiodination. There are other pathways of metabolism, such as ether bond cleavage [mainly of T4 in leukocytes (202)], deamination and decarboxylation of the alanine side chain (203,204), sulfoconjugation mediated by cytosolic phenol sulfotransferases in several tissues (205), and glucuronidation and O-methylation, which renders the products more hydrophilic and thereby facilitates their excretion by bile, feces, and urine (206).

Sulfoconjugation is the most important alternative iodothyronine metabolic pathway. The sulfotransferases of liver are normally involved in inactivation and detoxification reactions, with a preference for lipophilic substrates (207,208). Sulfation of iodothyronines facilitates rapid inner ring deiodination by D1 but not by D3. All iodothyronines except reverse T3, the preferred substrate of D1, are sulfated to some extent (205).

Iodothyronines with two iodine atoms at the phenolic ring are preferentially conjugated with glucuronic acid, whereas iodothyronines that contain only one iodine atom in the phenolic ring are sulfated with the following preference: 3′-T1 = 3,3′-T2 > T3 > rT3 > T4 (166,205,207,209,210,211). T3 can be sulfated by human cytosolic liver phenol sulfotransferase (EC 2.8.2.1), with Km values in the 100 mM range, considerably greater than that for D1 (212). Biliary and urinary excretion of iodothyronine sulfates is a minor route for thyroid hormone elimination in humans. Considerable amounts of iodothyronine sulfates are detectable in plasma and bile after inhibition of D1 by PTU in rats (205,213,214). Moreover, at a high substrate concentration (> 1 mMin vitro, metabolism proceeds mainly by sulfation, whereas at lower concentrations (< 0.1 mM), sulfation is the rate-limiting step; no sulfates accumulate because of the rapid deiodination of the sulfated iodothyronines. Thus far, the cellular capacity for conjugation seems to be unsaturable, in contrast to enzymic monodeiodination (29,166,209). In human hepatocarcinoma cells (HepG2 cells), inner ring deiodination of T3 is reduced, due to deficient T3 sulfation, which appears to be an obligatory step before deiodination of T3 (215).

Rats with selenium deficiency have high serum T3 sulfate concentrations and increased enterohepatic cycling of T3 sulfate, and the serum half-life of T3 is prolonged, as compared with normal rats (216). Under these conditions, sulfoconjugation might lead to greater availability of the active hormone. Sulfoconjugates of T4, T3, rT3, and 3,3′-T2 have been identified in human serum and amniotic fluid by specific radioimmunoassays (217,218,219,220,221). These conjugates are normal components of maternal and fetal serum and amniotic fluid. Their production and metabolism are regulated by the hormonal state, nonthyroidal illness, and drugs that inhibit deiodinase activity; sulfoconjugates of maternal origin may contribute to fetal thyroid economy.

PHYSIOLOGIC ROLES OF THE SELENODEIODINASES

D2 and Regulation of Thyrotropin Secretion

The main secretory product of the thyroid gland is T4, and serum T4 is a more important regulator of TSH secretion than serum T3. Because T3 is the biologically active thyroid hormone, T4 sensing must be preceded by its conversion to T3. The first evidence that there was a PTU-insensitive pathway for T4 to T3 conversion was that T4 very rapidly (within 30 minutes) inhibited TSH secretion in rats with hypothyroidism, and the inhibitory effect was not blocked by PTU (24,222,223,224). Subsequent studies in which combinations of 125I-T4 and 131I-T3 were injected revealed 125I-T3 bound to thyroid hormone receptors in the nuclei of pituitary cells within 15 minutes after injection of 125I-T4. This could not be explained by accumulation of 125I-T3 from serum, and was not inhibited by pretreatment with PTU (21,24). Pretreatment with iopanoic acid blocked both the generation of pituitary nuclear 125I-T3 and the biologic effect of T4 on TSH release (225).

The presence of D2 can account for the requirement for physiologic concentrations of both T4 and T3 for normal secretion of TSH. This requirement can account for the increase in TSH secretion that occurs in the early stages of iodine deficiency, when only T4, but not T3, production is decreased (226). Furthermore, normal serum concentrations of both T3 and T4 are required to suppress TRH mRNA production in the paraventricular nucleus of the hypothalamus and to normalize serum TSH concentrations in thyroidectomized rats (227,228,229). However, no D2 activity is present in this region of the hypothalamus; it is instead concentrated in the arcuate nucleus and median eminence (112,230). Subsequent in situ hybridization studies revealed that D2 is localized in the tanycytes (111,113,114). These specialized ependymal cells have their cell bodies in the inferior portion of the third ventricle, and they are probably where T4 is converted to T3, providing T3 that is released into the hypothalamic-pituitary portal system and carried to the thyrotrophs of the pituitary to regulate TSH secretion.

Triiodothyronine Homeostasis

The thyroid secretes T4 and T3 in a proportion determined by the T4/T3 ratio in thyroglobulin (15/1 in humans), as modified by the minimal thyroidal conversion of T4 to T3 (231). Thus, the prohormone T4 is the major secreted iodothyronine in iodine-sufficient subjects; the molar ratio of secreted T4 to T3 is about 11 to 1 due to intrathyroidal T4 to T3 conversion via D1 and D2 (116,232,233). As noted above, most T3 production occurs in various extrathyroidal tissues via 5′-deiodination of T4 catalyzed by D1 and D2. The serum concentrations of free T4 and T3 are constant, but the concentrations of T4 and T3 in cells vary according the amounts of each hormone that are transported or diffuse into the cells, and the type and activity of the deiodinases in the cells. These deiodinases can increase (D2) or decrease (D3) the intracellular concentrations of T3 and consequently the nuclear content of thyroid receptor-T3 complexes independently of the serum T4 and T3 concentrations. As a result, the impact of T4 and T3 in serum on thyroid hormone action varies in different tissues. In liver and kidney, for example, the saturation of the thyroid receptors is approximately 50%, whereas in the central nervous system it is close to 95%. Lastly, tissue T3 concentrations change throughout development, partially as a result of changes in the activity of D2 and D3. The deiodinases also modulate the thyroid status of individual tissues in response to iodine deficiency, hypothyroidism, and thyrotoxicosis. Cells lacking the capacity to alter the rate of deiodination of T4 and T3 are the most affected, because their thyroid status will be determined primarily by the serum free T3 concentration. On the other hand, in cells expressing D2 or D3, changes in the activity of these enzymes mitigate fluctuations in serum free T4 and T3concentrations, constituting a potent mechanism for maintaining thyroid homeostasis.

The relative contributions of the two sources of T3, thyroid secretion and extrathyroidal deiodination of T4, can be quantified by determining the T4 to T3conversion rate, which is, on average, about 35% to 40% in normal humans (60). Hence, with a normal T4 production rate of approximately 110 nmol/day (85 µg/day), approximately 40 nmol (25 µg) of T3 are produced by peripheral deiodination of T3 and the remaining 10 nmol (6 µg) are released by the thyroid (Fig. 7.8).

FIGURE 7.8. Daily triiodothyronine (T3) production in humans and rats. The dotted lines in the cylinder representing human extrathyroidal production reflect the uncertainty about the contributions of type 1 deiodinase (D1) and type 2 deiodinase (D2) to this pool. Values given are based on studies cited in the text. Estimates for rats are normalized to 100 g body weight. To convert nmol to µg, multiply by 0.65, and to convert pmol to µg, multiply by 0.00065.

Both D1 and D2 contribute to extrathyroidal production of T3, but assessing the contribution of the two deiodinases in vivo is difficult. In patients with primary hypothyroidism receiving constant doses of T4, administration of PTU in a dose of 1000 mg daily for 7 to 8 days caused a 20% to 30% decrease in serum T3concentrations (234,235). In another study, conversion of radiolabeled T4 to T3 in serum was not reduced in patients given 1200 mg/day of PTU (236). The results of these studies suggest that D1-catalyzed T3 production is not a major component of extrathyroidal T3 production in normal humans. However, in patients with thyrotoxicosis, the contribution from D1 is higher, about 50% (50), due to an increase in D1 activity. Fractional conversion of T4 to T3 is increased when serum T4 concentrations are low, indicative of increased D2 activity, because D1-catalyzed T3 production is decreased (50,237,238,239). It is difficult to define which compartments or tissues contribute the most to extrathyroidal T3 production in humans (240); depending on the assumptions used, one can obtain estimates suggesting that as much as 81% or as little as 15% of T3 derives from rapidly equilibrating (D1-containing) tissues, with the remainder coming from slowly equilibrating (D2-containing) tissues.

The relative contributions of the D1 and D2 pathways to whole-body T3 production can be assessed more accurately in rats. In normal rats treated chronically with high doses of PTU to inhibit D1 activity, the T4 to T3 conversion rate is reduced by 50% (241). Accordingly, in T4-treated thyroidectomized rats, treatment with PTU results in a 50% decrease in serum T3 concentrations (223). The results of compartmental analyses of T4 to T3 conversion rates in rats are similar, assuming that T4 to T3 conversion in the rapidly equilibrating pool occurs via D1 and more delayed conversion via D2 (242). Taken together, these data indicate that D1 catalyzes about half of the daily extrathyroidal T3 production from T4 in rats. This estimate of 50% in rats is clearly higher than the estimate of approximately 25% in humans from the above-mentioned PTU studies (Fig. 7.8).

There are several implications of the above calculations. Based on the data available, D2-catalyzed 5′-deiodination of T4 appears to be a more important source of T3 than does D1-catalyzed 5′-deiodination of T4 in humans. Until recently, D2 activity was believed to be restricted mainly to the central nervous system and pituitary. The identification of D2 mRNA and activity in skeletal muscle and heart suggests a more important role for D2 in T3 production (26,243). The fact that only approximately 20% of serum T3 comes from thyroidal secretion, as opposed to about 40% in rats, also indicates a need for more widespread T4 to T3 conversion in humans.

Intracellular Triiodothyronine Homeostasis

Serum T3 equilibrates rapidly with most tissues. Both T4 and T3 are carried across the plasma membrane by stereospecific energy-dependent transporters (244,245), and they may also diffuse into cells. At equilibrium, one can estimate the nuclear T3 derived from serum T3 from the ratio of nuclear to serum T3 and the serum T3 concentration. Measurements of the maximum binding capacity of the thyroid receptors for T3 allow calculation of the degree of saturation of the receptors, which is normally 40% to 50% in most tissues (246). Thus, changes in serum T3 concentrations during thyrotoxicosis or hypothyroidism are mirrored by changes in the occupancy of the thyroid receptors in those tissues, and it is the latter that determines the intensity of the biologic actions of thyroid hormone. However, in selected tissues, especially pituitary gland, brain, and brown adipose tissue, additional T3 is provided by intracellular T4 to T3 conversion (24,247). This has been termed T3 (T4) to differentiate it from T3 (T3), the nuclear T3 derived directly from serum T3. These tissues contain D2, and in them the T3 generated by D2-catalyzed T4 deiodination supplements that from serum as though it were derived from a kinetically different pool. As a result, thyroid receptor occupancy by T3 is much higher (70%–90%), and 50% to 80% of this receptor-bound T3 is T3 (T4) (21,23,248).

These differences have been confirmed using constant infusions of radiolabeled T3 and T4 (249,250,251,252) and direct quantitation of nuclear T3 (253). Other tissues (e.g., liver, kidney) in which serum T3 is the only source of nuclear T3 contain mostly D1. As discussed above, confocal microscopic studies of transiently expressed D2 suggest that it is located in the endoplasmic reticulum in the perinuclear region, and T3 formed in this region might have preferential access to the nucleus. D1, however, is distributed in the periphery of the cell, typical of a plasma membrane protein. The rapid exit of T3 from D1-containing tissues and its retention in D2-containing tissues explain the three- to four-fold higher ratio of nuclear to cytoplasmic free T3 in brain than in liver, kidney, or heart (254). The consequence of the presence of D2 is that the impact of changes in T4 production on cellular T3 content can be dampened at a prereceptor level by compensatory alterations in D2 activity.

Iodine Deficiency and Hypothyroidism

Iodine availability can be rate limiting for thyroidal T4 and T3 production, and multiple thyroidal and extrathyroidal mechanisms have evolved to mitigate the consequences of iodine deficiency (see the section on thyroid iodine transport in Chapter 4 and the section on iodine deficiency in Chapter 11) (255). Accordingly, in iodine-deficient rats, growth, O2 consumption, and thermal homeostasis are similar to those of normal rats, despite approximately 10-fold higher serum TSH concentrations and very low serum T4 concentrations (256,257). However, if iodine deficiency is severe and prolonged, signs of hypothyroidism do eventually develop, with reduced O2 consumption and reduced activity of T3-dependent enzymes (258,259,260). It is difficult to distinguish between compensated iodine deficiency and hypothyroidism, except by measurements of T3 actions, because serum TSH concentrations are high at all stages.

Iodine deficiency results in a series of physiologic adaptations in the hypothalamic–pituitary–thyroid axis, similar to those that occur in hypothyroidism, which maintain serum and tissue T3 concentrations in the normal range, delaying the onset of hypothyroidism. The earliest thyroidal adaptation is a decrease in diiodotyrosine formation, with a consequent decrease in thyroidal T4 synthesis, whereas thyroidal T3 synthesis is maintained (226,261). TSH se cretion increases as serum T4 concentrations decrease, increasing thyroidal iodide transport and the subsequent steps of thyroid hormone production (226,261). This stimulation helps to maintain thyroid hormone production, but with continuing iodine deficiency the ratio of diiodotyrosine to monoiodotyrosine in thyroglobulin decreases by approximately 3-fold and the ratio of T4 to T3 decreases by approximately 25-fold; the latter is due to a decrease in thyroidal T4content, not to an increase in T3 content. The ratio of T4 to T3 in the serum of iodine- deficient subjects is also low, due to hypothyroxinemia, not an increase in serum T3 concentrations. Thus, the hallmarks of iodine deficiency, and the early phase of primary hypothyroidism, are low serum T4, normal serum T3, and high serum TSH concentrations (261,262,263).

The extrathyroidal adaptations to iodine deficiency or primary hypothyroidism are more complex and involve a high degree of tissue specificity. As noted above, the overall fractional conversion of T4 to T3 is increased in patients with hypothyroidism (237). This increase is due to an increase in D2 activity, and its effect is to maintain extrathyroidal T3 production (26,96,243). In rats, the fractional T4 to T3 conversion rate is not substantially changed by hypothyroidism, but extrathyroidal T3 production shifts from being relatively PTU sensitive (~50%) to being PTU insensitive (264), indicating that the relative contribution of D2-catalyzed T4 conversion to T3 increases substantially.

In tissues that express D2, the activity of this enzyme is increased during iodine deficiency or hypothyroidism, thus increasing the local fractional conversion of T4 to T3, and mitigating the decrease in serum T4 concentrations (22,73,265,266,267). This has been particularly well documented for the brain; because of the negative regulation of Dio2 gene transcription by thyroid hormone (268), D2 mRNA increases in iodine-deficient animals in all regions of the brain that contain D2, and D2 activity increases even more (267), similar to what occurs in hypothyroid rats (134). This increase in D2 activity is explained by the hypothyroxinemia of iodine deficiency per se acting at a posttranslational level, as noted above in the sections on regulation of D2 synthesis and degradation.

In hypothyroidism, there is not only an increase in fractional conversion of T4 to T3, but the clearance of T3 from the brain also is reduced. This is because D3 is a T3-dependent gene, and its activity correlates with thyroid status. In both fetal and adult rats with iodine deficiency, overall brain D3 activity decreases by 50% (73,164,174,269,270). However, in specific brain regions, such as the cerebral cortex, hippocampus, and cerebellum, D3 activity decreases by 80% to 90% (73). The consequences of the decrease in D3 activity are twofold. First, the residence time of T3 within the tissue is prolonged (271). Second, because T4 is also a substrate for D3, relatively more of it will be available within the tissue for conversion to T3 by D2. Particularly in tissues like the brain, in which the exchange of T3 with serum is slow and most of the T3 is generated in situ, it is likely that fluctuations in the rate of T3 degradation have a greater influence on tissue levels of T3 than in tissues in which cellular T3 is in more rapid equilibration with serum, such as the liver and kidney (272).

The increased fractional production of T3 from T4 by D2 combined with the prolonged residence time of T3 mitigates the effects of severe iodine deficiency and hypothyroidism. This has been demonstrated in mild to moderate hypothyroidism using tracer studies (273). These pre dictions were confirmed directly by measuring thyroid hormone concentrations in various regions of the CNS in iodine-deficient rats (265). As expected, tissue T4 concentrations were markedly decreased, whereas tissue T3 concentrations were reduced by only 50%. This illustrates the effectiveness of these compensatory mechanisms.

Embryonic Development

Thyroid hormone is critically important for the coordination of developmental processes in all vertebrate species. During embryogenesis, thyroid hormone acts primarily to promote differentiation and thus attenuate proliferation. As a result, either insufficient levels of T3 or premature exposure of the embryo to high T3 concentrations result in abnormal development (274). As an example, exposure of neonatal rats to excess thyroid hormone causes accelerated morphogenesis of pyramidal neurons and their dendritic spines in the cerebrum and a persistent reduction in total number of neurons (275). The best characterized action by which thyroid hormone influences developmental processes is via changes in gene expression initiated by the binding of T3 to its receptors (276,277). During development in experimental animals, two deiodinases (D3 and D2) exert the major control of T3 concentrations (43). As mentioned earlier, fetal serum T3 concentrations are very low, and during early development D3 is the predominant deiodinase expressed in most rat tissues, and its activity in these tissues is much higher than in adult rats. In human fetuses, D3 is also present in the liver, and its content decreases near the end of gestation (167). This pattern suggests that D3 plays a major role in preventing premature exposure of fetal tissues to T3.

Conversely, D2 is expressed in most mammalian tissues for a limited period of time during development. This suggests that there is a tissue-specific program of T3-dependent differentiation. This has been found during tadpole metamorphosis, and it likely occurs during neuronal and glial maturation and cochlear maturation in rats (39,110,278,279,280,281). Finally, D1 activity is generally lower during fetal development than at later stages of life (282). This also would result in lower serum T3 concentrations at that time of life.

Deiodinases in Mammalian Development

Precisely timed D2 expression is fundamental during critical periods of mammalian development. In rat brain, D2 increases rapidly after birth, reaching its highest level around day 28. It then declines, reaching adult levels by day 50 (165). The cochlea is among the organs most sensitive to thyroid deficiency, as is evident from the deafness that is associated with endemic cretinism. In rats, complete cochlear maturation and the onset of auditory function require the presence of T3 between the late embryonic stage and the second postnatal week. So far, little is known about the mechanisms that control this temporal regulation. Analysis of cochlear homogenates from 2- to 8-day old rat pups revealed a striking peak of D2 activity peak at about 7 days, followed by an abrupt decline by day 10, a few days before the onset of hearing (110). Relative to serum, cochlear tissue has a high ratio of T3 to T4, supporting a role for D2 in increasing local T3 concentrations. D2 mRNA is localized in connective tissue near the region where dendritic and axonal projections of the cochlear nerve connect with the hair cells. D2 expression is complementary to, rather than coincident with, that of the βsubtype of the thyroid receptor, suggesting a paracrine rather than endocrine mode of signaling in cochlear tissue. D2-deficient mice have defective auditory function, retarded differentiation of the cochlear inner sulcus and sensory epithelium, and deformity of the tectorial membrane. The similarity of this phenotype to that caused by thyroid receptor mutations suggests that D2 controls the T3 signal that activates thyroid receptors in the cochlea (283).

Maternal-Fetal Physiology

In humans, the capacity to synthesize thyroid hormones does not appear until 10 to 12 weeks of gestation (see Chapter 74). However, human fetuses have thyroid receptors containing T3 at an earlier time (284); this T3 must come from the maternal circulation, via the placenta and the coelomic and amniotic fluids (285). At 6 to 12 weeks' gestation, the average total T4 concentrations are 11.3, 0.07, and 0.002 µg/dL (146, 0.96, and 0.02 nM), respectively, in maternal serum, coelomic fluid, and amniotic fluid, suggesting a marked gradient of T4 from mother to fetus. The gradient for reverse T3 is in the opposite direction, its concentrations being 3.8 and 15 times higher in the coelomic fluid and amniotic fluid, respectively, as compared with maternal serum (285). Also during the second and third trimesters, there are marked maternal-to-fetal gradients of free T4 and T3 (286,287).

D2 and D3 activity appear in fetal tissues at midgestation, whereas D1 is not evident until later (44). Accordingly, fetal serum T3 concentrations are very low before 30 weeks; thereafter they increase slightly, due to an increase in D1 activity in fetal tissues. The serum concentrations of sulfated iodothyronine concentrations are high in fetuses; although T3 sulfate does not bind to thyroid receptors, local desulfation, if it occurred, would provide a local source of T3 in fetal tissues (217,220,288,289). The pattern of circulating iodothyronines in the fetus, which is characterized by low serum T3 and high reverse T3concentrations, is due to the combination of high D3 activity and low D1 activity in fetal tissues throughout most of gestation.

Triiodothyronine Modulation of Placental Thyroid Hormone Transfer

The placenta is the pathway for maternal-fetal thyroid hormone transfer, and therefore can be an important determinant of the thyroid status of the fetus. Placental D3 activity increases with gestational age in rats and humans (176,192,290,291). In the first trimester, when the placenta and the surface area available for transfer are small, the activity of D3 per unit of placental tissue is high. At term, the activity per unit is lower, but because the placenta is much larger total placental D3 activity is higher. In rats, unlike humans, placental D3 activity increases about twofold from embryonic day 14 until day 16 or 17, after which it decreases (176,292,293). As mentioned above, placenta also contains D2; however, at all gestational ages, placental D3 activity is approximately 200-fold higher than is that of D2. In placental tissue, there is no direct correlation between D2 activity and mRNA levels, and although D3 activity is always higher than D2 activity, their mRNA levels are comparable (291).

The cellular localization of D2 and D3 in placental tissue also differs. D2 activity is higher in the chorionic and decidual membranes of the placenta than in the amniotic membranes, whereas D3 is found mostly in trophoblasts (194). However, given the low levels of D2 at all gestational ages, fluctuations in D2 activity are not likely to have a major effect on fetal serum T3 concentrations.

The physiologic consequence of the high placental D3 activity is clear. Placental tissue actively deiodinates T4 to reverse T3 and T3 to 3,3′-diiodothyronine, some of which is further deiodinated to 3′-monoiodothyronine (191). In isolated perfused human placental lobules, little of the T4 added to the maternal side appears on the fetal side. In contrast, reverse T3 concentrations rise progressively on both sides. Addition of the deiodinase inhibitor iopanoic acid to the maternal perfusate results in an increase in T4 and a decrease in reverse T3 on the fetal side, providing direct evidence that human placental D3 is a major factor controlling transfer of maternal T4 to the fetus (294).

The uterus of pregnant rats also contains high levels of D3 activity, initially in decidual cells and later in the single-cell layer of the epithelium (195). D3 activity is highest at the implantation site, and the activity there is almost double the highest value found in the placenta. These changes precede the appearance of thyroid receptor mRNA (295). Throughout gestation, D3 activity remains higher in the uterus than in the placenta, and is 10 times higher than in the entire fetus. D3 activity has also been detected in amniotic fluid (195). High uterine and placental D3 activity also protects against high T3concentrations, which can induce structural abnormalities in the cephalic and branchial arches (296).

The presence of D3 in the endometrium and placenta villae as well as every epithelial surface of the human fetus constitutes a barrier to inappropriate transfer of maternal T4 and T3 to the fetus (197). This barrier is so potent that instillation of 700 µg of T4 into amniotic fluid at term results in little increase in umbilical cord serum T3 concentrations of infants born 24 hours later (297). The high level of D3 activity in the uterus and placenta contributes to the need for more T4 in women with hypothyroidism when they are pregnant (298).

Paradoxically, despite inactivation of T4 and T3 by the uterus and placenta, neonates with congenital hypothyroidism often have little evidence of hypothyroidism at birth. Among infants with a complete defect in thyroid iodide organification, cord serum T4 concentrations are 20% to 50% of those in normal infants, indicating there is substantial transplacental passage of T4. The concentrations decrease rapidly after birth in these infants (299). These results indicate that a steep maternal-fetal gradient of T4 overcomes the placental barrier, permitting maternal T4 to enter the fetal circulation.

THE DEIODINASES IN HUMAN PATHOPHYSIOLOGY

Nonthyroidal Illness

Extrathyroidal conversion of T4 to T3 is decreased in patients with virtually all nonthyroidal illness, as discussed above and in the section on nonthyroidal illness in Chapter 11.

Consumptive Hypothyroidism

As noted above, high levels of D3 are expressed in hemangiomas. If these tumors are sufficiently large, the rate of T4 and T3 deiodination can exceed thyroidal production of the hormones, even in the presence of intense TSH stimulation. The first patient documented to have this condition was a 3-month-old infant who had severe hypothyroidism, with high serum TSH, undetectable T4 and T3, and high reverse T3 concentrations. To reverse the hypothyroidism rapidly, the infant was given T3 and T4 intravenously in doses sufficient to reduce TSH secretion to normal. The respective daily doses were 96 µg of T3 and 50 µg of T4 (162), doses that would cause overt thyrotoxirosis in not only normal infants but also normal adults (300). Remarkably, even during the infusions, serum T3 concentrations barely reached the normal range, serum T4 was never detectable, and serum reverse T3 concentrations were very high, providing direct evidence of excessive inner ring deiodination. D3 activity was subsequently identified in the infant's hemangioma at levels eight times that in the placenta, and in situ hybridization localized the D3 mRNA to hemangioma cells. Other patients with this unique syndrome have now been reported (301).

The relationship between infantile hemangiomas and D3 expression is important because it identifies a cause of hypothyroidism at a critical age for neurologic development. Although large hepatic hemangiomas can be fatal, a substantial fraction of these infants survive with therapy and the natural propensity of these tumors to regress. If hypothyroid, these patients may require high doses of T4 in addition to therapy directed at their hemangiomas.

D3 Expression and the Requirement for Thyroxine in Normal Pregnancy

Most pregnant women with hypothyroidism need an increase in T4 dose of approximately 40% to maintain normal serum TSH concentrations (299,302). The need for an increase begins early in pregnancy and persists until delivery (303), and its existence indicates that T4 and T3 production must increase in normal pregnant women. The causes of this need for more T4 and T3 include an increase in uterine D3 activity and the presence of placental D3 activity, transfer of T4and T3 to the fetus, and the estrogen-induced increase in serum thyroxine-binding globulin concentrations.

Overexpression of Deiodinases and Excess Triiodothyronine Production

In patients with thyrotoxicosis, there is a disproportionate increase in the production rate of T3 and serum T3 concentrations, as compared with those of T4, by a factor of two (50). The human Dio1 gene promoter is T3-responsive, and D1 mRNA levels are increased in thyroid tissue and mononuclear leukocytes in patients with thyrotoxicosis (42,304,305). In these patients, PTU—which blocks D1, but not D2, activity—results in a more rapid acute decrease in serum T3concentrations than does methimazole, which has no effect of either D1 or D2 activity (50). These results indicate that D1-catalyzed T4 to T3 conversion is increased in patients with thyrotoxicosis (234,235,236). This has led to the recommendation that patients with severe thyrotoxicosis be treated with large doses of PTU or other drugs that block T4 conversion to T3 (see Chapter 45) (306,307,308,309).

In patients with Graves' thyrotoxicosis, thyroidal D2 mRNA is increased despite high serum T4 and T3 concentrations (116). This is presumably due to the effect of TSH receptor–stimulating antibodies to activate the cyclic AMP-dependent Dio2 gene, which overwhelms the negative transcriptional effect of T3 on this gene. Therefore, some of the relative excess of T3 production in patients with thyrotoxicosis is caused by an increase in thyroidal D2 activity (116).

Another consequence of thyroid overexpression of D2 is illustrated by three athyreotic patients with widespread follicular carcinoma of the thyroid in whom the ratio of T3 to T4 in serum was persistently high, in the absence of autonomous production of T3 by the tumor. In one patient, the tumor contained a high level of D2 activity, and resection of the tumor resulted in a normal T4 to T3 ratio in serum. In two other patients, treatment with T4 in doses sufficient to suppress TSH secretion was associated with high normal serum T3 and subnormal free T4 values (310). These findings indicate that increases in T4 conversion to T3, presumably caused by increased D2 activity, can alter serum T4 and T3 concentrations.

CONCLUSION

Many tissues contain deiodinase activity, and it is now clear that these enzymes are important determinants of thyroid hormone homeostasis in both health and disease. Two deiodinases catalyze the intracellular conversion of T4 to T3, and a third catalyzes intracellular conversion of T4 and T3 to biologically inactive products. Variations in the activity of these enzymes, as a result of thyroid and other disorders, can affect thyroid hormone availability and therefore thyroid hormone action, in many tissues.

ACKNOWLEDGMENT

This work was supported by National Institutes of Health Grants DK36256, DK44128, and DK58538.

REFERENCES

1. Bianco AC, Salvatore D, Gereben B, et al. Biochemistry, cellular and molecular biology and physiological roles of the iodothyronine selenodeiodinases.Endocr Rev 2002;23:38.

2. Berry MJ, Kieffer JD, Harney JW, et al. Selenocysteine confers the biochemical properties of the type I iodothyronine deiodinase. J Biol Chem 1991;266:14155.

3. Croteau W, Whittemore SL, Schneider MJ, et al. Cloning and expression of a cDNA for a mammalian type III iodothyronine deiodinase. J Biol Chem 1995;270:16569.

4. Buettner C, Harney JW, Larsen PR. The role of selenocysteine 133 in catalysis by the human type 2 iodothyronine deiodinase. Endocrinology 2000;141:4606.

5. Callebaut I, Curcio-Morelli C, Mornon JP, et al. The iodothyronine selenodeiodinases are thioredoxin-fold family proteins containing a glycoside hydrolase-clan GH-A-like structure. J Biol Chem 2003;278:36887.

6. Martin JL. Thioredoxin—a fold for all reasons. Structure 1995; 3:245.

7. Coutinho PM, Henrissat B. Carbohydrate-active enzymes. http://afmb.cnrs-mrs.fr/CAZY, accessed 9 August 2004.

8. Berry MJ, Banu L, Larsen PR. Type I iodothyronine deiodinase is a selenocysteine-containing enzyme. Nature 1991;349:438.

9. Berry MJ, Banu L, Chen YY, et al. Recognition of UGA as a selenocysteine codon in type I deiodinase requires sequences in the 3′ untranslated region. Nature 1991;353:273.

10. Leonard JL, Visser TJ, Leonard DM. Characterization of the subunit structure of the catalytically active type I iodothyronine deiodinase. J Biol Chem 2000;276:2600.

11. Curcio-Morelli C, Gereben B, Zavacki AM, et al. In vivo dimerization of types 1, 2, and 3 iodothyronine selenodeiodinases. Endocrinology 2003;144:3438.

12. Leonard JL, Rosenberg IN. Solubilization of a phospholipid-requiring enzyme, iodothyronine 5′-deiodinase, from rat kidney membranes. Biochim Biophys Acta 1981;659:205.

13. Safran M, Leonard JL. Comparison of the physicochemical properties of type I and type II iodothyronine 5′-deiodinase. J Biol Chem 1991;266:3233.

14. Toyoda N, Berry MJ, Harney JW, et al. Topological analysis of the integral membrane protein, type 1 iodothyronine deiodinase (D1). J Biol Chem 1995;270:12310.

15. Baqui MM, Gereben B, Harney JW, et al. Distinct subcellular localization of transiently expressed types 1 and 2 iodothyronine deiodinases as determined by immunofluorescence confocal microscopy. Endocrinology 2000;141:4309.

16. Leonard JL, Rosenberg IN. Subcellular distribution of thyroxine 5′-deiodinase in the rat kidney: a plasma membrane location. Endocrinology 1978;103:274.

17. Kohrle J, Rasmussen UB, Rokos H, et al. Selective affinity labeling of a 27-kd integral membrane protein in rat liver and kidney with N -bromoacetyl derivatives of L-thyroxine and 3,5,3′-triiodo-L-thyronine. J Biol Chem 1990;265:6146.

18. Prabakaran D, Ahima RS, Harney JW, et al. Polarized targeting of epithelial cell proteins in thyrocytes and MDCK cells. J Cell Sci 1999;112:1247.

19. Baqui MM, Botero D, Gereben B, et al. Human type 3 iodothyronine selenodeiodinase is located in the plasma membrane and undergoes rapid internalization to endosomes. J Biol Chem 2003;278:1206.

20. Curcio C, Baqui MMA, Salvatore D, et al. The human type 2 iodothyronine deiodinase is a selenoprotein highly expressed in a mesothelioma cell line. J Biol Chem 2001;276:30183.

21. Silva JE, Larsen PR. Contributions of plasma triiodothyronine and local thyroxine monodeiodination to triiodothyronine to nuclear triiodothyronine receptor saturation in pituitary, liver, and kidney of hypothyroid rats: further evidence relating saturation of pituitary nuclear triiodothyronine receptors and the acute inhibition of thyroid-stimulating hormone release. J Clin Invest 1978;61:1247.

22. Larsen PR, Silva JE, Kaplan MM. Relationships between circulating and intracellular thyroid hormones: physiological and clinical implications. Endocr Rev 1981;2:87.

23. Silva JE, Dick TE, Larsen PR. The contribution of local tissue thyroxine monodeiodination to the nuclear 3,5,3′-triiodothyronine in pituitary, liver, and kidney of euthyroid rats. Endocrinology 1978;103:1196.

24. Silva JE, Larsen PR. Pituitary nuclear 3,5,3′-triiodothyronine and thyrotropin secretion: an explanation for the effect of thyroxine. Science 1977;198:617.

25. Leonard JL, Visser TJ. Biochemistry of deiodination. In: Hennemann G, ed. Thyroid hormone metabolism. New York: Marcel Dekker, 1986:189.

26. Salvatore D, Bartha T, Harney JW, et al. Molecular biological and biochemical characterization of the human type 2 selenodeiodinase. Endocrinology 1996;137:3308.

27. Kuiper GG, Klootwijk W, Visser TJ. Substitution of cysteine for a conserved alanine residue in the catalytic center of type II iodothyronine deiodinase alters interaction with reducing cofactor. Endocrinology 2002;143:1190.

28. Sanders JP, Van der Geyten S, Kaptein E, et al. Characterization of a propylthiouracil-insensitive type I iodothyronine de iodinase. Endocrinology 1997;138:5153.

29. Otten MH, Mol JA, Visser TJ. Sulfation preceding deiodination of iodothyronines in rat hepatocytes. Science 1983;221:81.

30. Berry MJ. Identification of essential histidine residues in rat type I iodothyronine deiodinase. J Biol Chem 1992;267:18055.

31. Kohrle J, Hesch RD. Biochemical characteristics of iodothyronine monodeiodination by rat liver microsomes: the interaction between iodothyronine substrate analogs and the ligand binding site of the iodothyronine deiodinase resembles that of the TBPA-iodothyronine ligand binding. Horm Metab Res 1984;14:42(suppl).

32. Fekkes D, Hennemann G, Visser TJ. Evidence for a single enzyme in rat liver catalyzing the deiodination of the tyrosyl and the phenolic ring of iodothyronines. Biochem J 1982;201:673.

33. Jakobs TC, Koehler MR, Schmutzler C, et al. Structure of the human type I iodothyronine 5′-deiodinase gene and localization to chromosome 1p32-p33. Genomics 1997;42:361.

34. Mandel SJ, Berry MJ, Kieffer JD, et al. 1992 Cloning and in vitro expression of the human selenoprotein, type I iodothyronine deiodinase. J Clin Endocrinol Metab 1992;75:1133.

35. Maia AL, Berry MJ, Sabbag R, et al. Structural and functional differences in the dio1 gene in mice with inherited type 1 deiodinase deficiency. Mol Endocrinol 1995;9:969.

36. Toyoda N, Harney JW, Berry MJ, et al. Identification of critical amino acids for 3,5,3′-triiodothyronine deiodination by human type 1 deiodinase based on comparative functional-structural analyses of the human, dog, and rat enzymes. J Biol Chem 1994; 269:20329.

37. Van der Geyten S, Sanders JP, Kaptein E, et al. Expression of chicken hepatic type I and type III iodothyronine deiodinases during embryonic development. Endocrinology 1997;138:5144.

38. Galton VA. Iodothyronine 5′-deiodinase activity in the amphibian Rana catesbeiana at different stages of the life cycle. Endocrinology 1988;122:1746.

39. Becker KB, Stephens KC, Davey JC, et al. The type 2 and type 3 iodothyronine deiodinases play important roles in coordinating development in Rana catesbeiana tadpoles. Endocrinology 1997;138:2989.

40. St. Germain DL, Galton VA. The deiodinase family of selenoproteins. Thyroid 1997;7:655.

41. Campos-Barros A, Hoell T, Musa A, et al. Phenolic and tyrosyl ring iodothyronine deiodination and thyroid hormone concentrations in the human central nervous system. J Clin Endo crinol Metab 1996;81:2179.

42. Nishikawa M, Toyoda N, Yonemoto T, et al. Quantitative measurements for type 1 deiodinase messenger ribonucleic acid in human peripheral blood mononuclear cells: mechanism of the preferential increase of T3 in hyperthyroid Graves' disease. Biochem Biophys Res Commun 1998;250:642.

43. Bates JM, St. Germain DL, Galton VA. Expression profiles of the three iodothyronine deiodinases, D1, D2, and D3, in the developing rat. Endocrinology 1999;140:844.

44. Burrow GN, Fisher DA, Larsen PR. Maternal and fetal thyroid function. N Engl J Med 1994;331:1072.

45. Berry MJ, Kates AL, Larsen PR. Thyroid hormone regulates type I deiodinase messenger RNA in rat liver. Mol Endocrinol 1990;4:743.

46. Toyoda N, Zavacki AM, Maia AL, et al. A novel retinoid X receptor-independent thyroid hormone response element is present in the human type 1 deiodinase gene. Mol Cell Biol 1995; 15:5100.

47. Jakobs TC, Schmutzler C, Meissner J, et al. The promoter of the human type I 5′-deiodinase gene-mapping of the transcription start site and identification of a DR+4 thyroid-hormone–responsive element. Eur J Biochem 1997;247:288.

48. Zhang C, Kim S, Harney JW, et al. Further characterization of thyroid hormone response elements in the human type 1 iodothyronine deiodinase gene. Endocrinology 1998;139: 1156.

49. Amma LL, Campos-Barros A, Wang Z, et al. Distinct tissue-specific roles for thyroid hormone receptors beta and alpha1 in regulation of type 1 deiodinase expression. Mol Endocrinol 2001;15:467.

50. Abuid J, Larsen PR. Triiodothyronine and thyroxine in hyperthyroidism: comparison of the acute changes during therapy with antithyroid agents. J Clin Invest 1974;54:201.

51. Chopra IJ, Sakane S, Chua Teco GN. A study of the serum concentration of tumor necrosis factor-α in thyroidal and nonthyroidal illnesses. J Clin Endocrinol Metab 1991;72: 1113.

52. Boelen A, Platvoet-Ter Schiphorst MC, Wiersinga WM. Association between serum interleukin-6 and serum 3,5,3′- triiodothyronine in nonthyroidal illness. J Clin Endocrinol Metab 1993;77:1695.

53. van der Poll T, Romijn JA, Wiersinga WM, et al. Tumor necrosis factor: a putative mediator of the sick euthyroid syndrome in man. J Clin Endocrinol Metab 1990;71:1567.

54. Pekary AE, Berg L, Santini F, et al. Cytokines modulate type I iodothyronine deiodinase mRNA levels and enzyme activity in FRTL-5 rat thyroid cells. Mol Cell Endocrinol 1994;101:R31.

55. Nagaya T, Fujieda M, Otsuka G, et al. A potential role of activated NF-kappa B in the pathogenesis of euthyroid sick syndrome. J Clin Invest 2000;106:393.

56. Yu J, Koenig RJ. Regulation of hepatocyte thyroxine 5′-deiodinase by T3 and nuclear receptor coactivators as a model of the sick euthyroid syndrome. J Biol Chem 2000;275:38296.

57. Portnay GI, O'Brien JT, Bush J, et al. The effect of starvation on the concentration and binding of thyroxine and triiodothyronine in serum and on the response to TRH. J Clin Endocrinol Metab 1974;39:191.

58. Burrows AW, Shakespear RA, Hesch RD, et al. Thyroid hormones in the elderly sick: “T4 euthyroidism.” BMJ 1975;4:437.

59. Kaplan MM, Larsen PR, Crantz FR, et al. Prevalence of abnormal thyroid function test results in patients with acute medical illnesses. Am J Med 1992;72:9.

60. Larsen PR, Davies TF, Hay ID. The thyroid gland. In: Wilson JD, Foster DW, Kronenberg HM, et al., eds. Williams textbook of endocrinology, 9th ed. Philadelphia: WB Saunders, 1998:389.

61. Peeters RP, Wouters PJ, Kaptein E, et al. Reduced activation and increased inactivation of thyroid hormone in tissues of critically ill patients. J Clin Endocrinol Metab 2003;88:3202.

62. Harris ARC, Fang SL, Vagenakis AG, et al. Effect of starvation, nutrient replacement, and hypothyroidism on in vitro hepatic T4 to T3 conversion in the rat. Metabolism 1978;27: 1680.

63. Harris ARC, Fang SL, Hinerfeld L, et al. The role of sulfhydryl groups on the impaired hepatic 3′,3,5-triiodothyronine generation from thyroxine in the hypothyroid, starved, fetal and neonatal rodent. J Clin Invest 1979;63:516.

64. Goodman MN, Larsen PR, Kaplan MM, et al. Starvation in the rat. II. Effect of age and obesity on protein sparing and fuel metabolism. Am J Physiol 1980;239:E277.

65. Kinlaw WB, Schwartz HL, Oppenheimer JH. Decreased serum triiodothyronine in starving rats is due primarily to diminished thyroidal secretion of thyroxine. J Clin Invest 1984;75:1238.

66. Safran M, Kohrle J, Braverman LE, et al. Effect of biological alterations of type I 5′deiodinase activity on affinity labeled membrane proteins in rat liver and kidney. Endocrinology 1990; 126:826.

67. Legradi G, Emerson CH, Ahima RS, et al. Leptin prevents fasting-induced suppression of prothyrotropin-releasing hormone messenger ribonucleic acid in neurons of the hypothalamic paraventricular nucleus. Endocrinology 1997;138:2569.

68. Vignati L, Finley RJ, Hagg S, et al. Protein conservation during prolonged fast: a function of triiodothyronine levels. Trans Assoc Am Physicians 1978;91:169.

69. Beckett GJ, Beddows SE, Morrice PC, et al. Inhibition of hepatic deiodination of thyroxine is caused by selenium deficiency in rats. Biochem J 1987;248:443.

70. Beckett GJ, MacDougal DA, Nicol F, et al. Inhibition of type I and II iodothyronine deiodinase activity in rat liver, kidney and brain produced by selenium deficiency. Biochem J 1989;259: 887.

71. Arthur JR, Nicol F, Beckett GJ. Hepatic iodothyronine 5′-deiodinase. The role of selenium. Biochem J 1990;272:537.

72. Behne D, Kyriakopoulos A, Meinhold H, et al. Identification of type I iodothyronine 5′-deiodinase as a selenoenzyme. Biochem Biophys Res Commun 1990;173:1143.

73. Meinhold H, Campos-Barros A, Behne D. Effects of selenium and iodine deficiency on iodothyronine deiodinases in brain, thyroid and peripheral tissue. Acta Med Aust 1992;19:8.

74. Meinhold H, Campos-Barros A, Walzog B, et al. Effects of selenium and iodine deficiency on type I, type II and type III iodothyronine deiodinases and circulating thyroid hormones in the rat. Exp Clin Endocrinol 1993;101:87.

75. DePalo D, Kinlaw WB, Zhao C, et al. Effect of selenium deficiency on type I 5′-deiodinase. J Biol Chem 1994;269:16223.

76. Calomme M, Vanderpas J, Francois B, et al. Effects of selenium supplementation on thyroid hormone metabolism in phenylketonuria subjects on a phenylalanine restricted diet. Biol Trace Elem Res 1995;47:349.

77. Jochum F, Terwolbeck K, Meinhold H, et al. Effects of a low selenium state in patients with phenylketonuria. Acta Paediatr 1997;86:775.

78. Lombeck I, Jochum F, Terwolbeck K. Selenium status in infants and children with phenylketonuria and in maternal phenylketonuria. Eur J Pediatr 1996;155 (suppl):140.

79. Kauf E, Dawczynski H, Jahreis G, et al. Sodium selenite therapy and thyroid-hormone status in cystic fibrosis and congenital hypothyroidism. Biol Trace Elem Res 1994;40:247.

80. Vanderpas JB, Contempre B, Duale NL, et al. Iodine and selenium deficiency associated with cretinism in northern Zaire. Am J Clin Nutr 1990;52:1087.

81. Goyens P, Golstein J, Nsombola B, et al. Selenium deficiency as a possible factor in the pathogenesis of myxoedematous endemic cretinism. Acta Endocrinol (Copenh) 1987;114:497.

82. Contempre B, Dumont JE, Ngo B, et al. Effect of selenium supplementation in hypothyroid subjects of an iodine and selenium deficient area: the possible danger of indiscriminate supplementation of iodine-deficient subjects with selenium. J Clin Endocrinol Metab 1991;73:213.

83. Contempre B, Duale NL, Dumont JE, et al. Effect of selenium supplementation on thyroid hormone metabolism in an iodine and selenium deficient population. Clin Endocrinol (Oxf) 1992; 36:579.

84. Steinsapir J, Harney J, Larsen PR.Type 2 iodothyronine deiodinase in rat pituitary tumor cells is inactivated in proteasomes. J Clin Invest 1998;102:1895.

85. Steinsapir J, Bianco AC, Buettner C, et al. Substrate-induced down-regulation of human type 2 deiodinase (hD2) is mediated through proteasomal degradation and requires interaction with the enzyme's active center. Endocrinology 2000;141:1127.

86. Gereben B, Goncalves C, Harney JW, et al. Selective proteolysis of human type 2 deiodinase: a novel ubiquitin-proteasomal mediated mechanism for regulation of hormone activation. Mol Endocrinol 200;14:1697.

87. Celi FS, Canettieri G, Mentuccia D, et al. Structural organization and chromosomal localization of the human type II deiodinase gene. Eur J Endocrinol 200;143:267.

88. Araki O, Murakami M, Morimura T, et al. Assignment of type II iodothyronine deiodinase gene (DIO2) to human chromosome band 14q24.2 & q24.3 by in situ hybridization. Cytogenet Cell Genet 1999;84:73.

89. Davey JC, Schneider MJ, Becker KB, et al. Cloning of a 5.8 kb cDNA for a mouse type 2 deiodinase. Endocrinology 1999; 140:1022.

90. Bartha T, Kim SW, Salvatore D, et al. Characterization of the 5′-flanking and 5′-untranslated regions of the cyclic adenosine 3′,5′-monophosphate-responsive human type 2 iodothyronine deiodinase gene. Endocrinology 2000;141:229.

91. Ohba K, Yoshioka T, Muraki T. Identification of two novel splicing variants of human type II iodothyronine deiodinase mRNA. Mol Cell Endocrinol 2001;172:169.

92. Song S, Adachi K, Katsuyama M, et al. Isolation and characterization of the 5′-upstream and untranslated regions of the mouse type II iodothyronine deiodinase gene. Mol Cell Endocrinol 2000;165:189.

93. Gereben B, Salvatore D, Harney JW, et al. The human, but not rat, dio2 gene is stimulated by thyroid transcription factor-1 (TTF-1). Mol Endocrinol 2001;15:112.

94. Buettner C, Harney JW, Larsen PR. The 3′-untranslated region of human type 2 iodothyronine deiodinase mRNA contains a functional selenocysteine insertion sequence element. J Biol Chem 1998;273:33374.

95. Gereben B, Bartha T, Tu HM, et al. Cloning and expression of the chicken type 2 iodothyronine 5′-deiodinase. J Biol Chem 1999;274:13768.

96. Croteau W, Davey JC, Galton VA, et al. Cloning of the mammalian type II iodothyronine deiodinase: a selenoprotein differentially expressed and regulated in human and rat brain and other tissues. J Clin Invest 1996;98:405.

97. Davey JC, Becker KB, Schneider MJ, et al. Cloning of a cDNA for the type II iodothyronine deiodinase. J Biol Chem 1995; 270:26786.

98. Valverde C, Croteau W, Lafleur GJ, Jr, et al. Cloning and expression of a 5′-iodothyronine deiodinase from the liver of Fundulus heteroclitus. Endocrinology 1997;138:642.

99. Salvatore D, Harney JW, Larsen PR. Mutation of the Secys residue 266 in human type 2 selenodeiodinase alters 75Se incorporation without affecting its biochemical properties. Bio chimie 1999;81:1.

100. Cheron RG, Kaplan MM, Larsen PR. Physiological and pharmacological influences on thyroxine to 3,5,3′-triiodothyronine conversion and nuclear 3,5,3′-triiodothyronine binding in rat anterior pituitary. J Clin Invest 1979;64:1402.

101. Leonard JL. Dibutyryl cAMP induction of type II 5′deiodinase activity in rat brain astrocytes in culture. Biochem Biophys Res Commun 1988;151:1164.

102. Visser TJ, Leonard JL, Kaplan MM, et al. Kinetic evidence suggesting two mechanisms for iodothyronine 5′-deiodination in rat cerebral cortex. Proc Natl Acad Sci USA 1982; 79:5080.

103. Crantz FR, Larsen PR. Rapid thyroxine to 3,5,3′-triiodothyronine conversion and nuclear 3,5,3′-triiodothyronine binding in rat cerebral cortex and cerebellum. J Clin Invest 19980;65: 935.

104. Silva JE, Larsen PR. Adrenergic activation of triiodothyronine production in brown adipose tissue. Nature 1983;305:712.

105. Molinero P, Osuna C, Guerrero JM. Type II thyroxine 5′-deiodinase in the rat thymus. J Endocrinol 195;146:105.

106. Song S, Sorimachi K, Adachi K, et al. Biochemical and molecular biological evidence for the presence of type II iodothyronine deiodinase in mouse mammary gland. Mol Cell Endocrinol 2000;160:173.

107. Kamiya Y, Murakami M, Araki O, et al. Pretranslational regulation of rhythmic type II iodothyronine deiodinase expression by beta-adrenergic mechanism in the rat pineal gland. Endocrinology 199;140:1272.

108. Galton VA, Martinez E, Hernandez A, et al. The type 2 iodothyronine deiodinase is expressed in the rat uterus and induced during pregnancy. Endocrinology 2001;142:2123.

109. Mizuma H, Murakami M, Mori M. Thyroid hormone activation in human vascular smooth muscle cells: expression of type II iodothyronine deiodinase. Circ Res 2001;88:313.

110. Campos-Barros A, Amma LL, Faris JS, et al. Type 2 iodothyronine deiodinase expression in the cochlea before the onset of hearing. Proc Natl Acad Sci USA 2000;97:1287.

111. Guadano-Ferraz A, Obregon MJ, St. Germain DL, et al. The type 2 iodothyronine deiodinase is expressed primarily in glial cells in the neonatal rat brain. Proc Natl Acad Sci USA 1997; 94:10391.

112. Riskind PN, Kolodny JM, Larsen PR. The regional hypothalamic distribution of type II 5′-monodeiodinase in euthyroid and hypothyroid rats. Brain Res 1987;420:194.

113. Tu HM, Kim SW, Salvatore D, et al. Regional distribution of type 2 thyroxine deiodinase messenger ribonucleic acid in rat hypothalamus and pituitary and its regulation by thyroid hormone. Endocrinology 1997;138:3359–3368

114. Fekete C, Mihaly E, Herscovici S, et al. DARPP-32 and CREB are present in type 2 iodothyronine deiodinase-producing tanycytes: implications for the regulation of type 2 deiodinase activity. Brain Res 2000;862:154.

115. Diano S, Naftolin F, Goglia F, et al. Monosynaptic pathway between the arcuate nucleus expressing glial type II iodothyronine 5′-deiodinase mRNA and the median eminence projective TRH cells of the rat paraventricular nucleus. J Neuroendocrinol 1998;10:731.

116. Salvatore D, Tu H, Harney JW, et al. 1996 Type 2 iodothyronine deiodinase is highly expressed in human thyroid. J Clin Invest 1996;98:962.

117. Murakami M, Araki O, Hosoi Y, et al. Expression and regulation of type II iodothyronine deiodinase in human thyroid gland. Endocrinology 2001;142:2961.

118. Imai Y, Toyoda N, Maeda A, et al. Type 2 iodothyronine deiodinase expression is upregulated by protein kinase A–dependent pathway and is downregulated by the protein kinase C–dependent pathway in cultured human thyroid cells. Thyroid 2001; 11:899.

119. Itagaki Y, Yoshida K, Ikede H, et al. Thyroxine 5′-deiodinase in human anterior pituitary tumors. J Clin Endocrinol Metab 1990;71:340.

120. Murakami M, Araki O, Morimura T, et al. Expression of type II iodothyronine deiodinase in brain tumors. J Clin Endocrinol Metab 2000;85:4403.

121. Kaplan MM, Pan C, Gordon PR, et al. Human epidermal keratinocytes in culture convert thyroxine to 3,5,3′-triiodothyronine by type II iodothyronine deiodination: a novel endocrine function of the skin. J Clin Endocrinol Metab 1988;66:815.

122. Salvatore D, Bartha T, Larsen PR. The guanosine monophosphate reductase gene is conserved in rats and its expression increases rapidly in brown adipose tissue during cold exposure. J Biol Chem 1998;273:31092.

123. Mills I, Raasmaja A, Moolten N, et al. Effect of thyroid status on catecholamine stimulation of thyroxine 5′-deiodinase in brown adipocytes. Am J Physiol 1989;256:E74.

124. Raasmaja A, Larsen PR. Alpha 1- and beta-adrenergic agents cause synergistic stimulation of the iodothyronine deiodinase in rat brown adipocytes. Endocrinology 1989;125:2502.

125. Noronha M, Raasmaja A, Moolten N, et al. Triiodothyronine causes rapid reversal of alpha 1/cyclic adenosine monophosphate synergism on brown adipocyte respiration and type II deiodinase activity. Metabolism 1991;40:1327.

126. Silva JE, Larsen PR. Hormonal regulation of iodothyronine 5′-deiodinase in rat brown adipose tissue. Am J Physiol 1986;251: E639.

127. Silva JE, Larsen PR. Interrelationships among thyroxine, growth hormone, and the sympathetic nervous system in the regulation of 5′-iodothyronine deiodinase in rat brown adipose tissue. J Clin Invest 1986;77:1214.

128. Pallud S, Lennon AM, Ramauge M, et al. Expression of the type II iodothyronine deiodinase in cultured rat astrocytes is selenium-dependent. J Biol Chem 1997;272:18104.

129. Gondou A, Toyoda N, Nishikawa M, et al. Induction of type 2 deiodinase activity by cyclic guanosine 3′,5′-monophosphate in cultured rat glial cells. Thyroid 1998;8:615.

130. Gondou A, Toyoda N, Nishikawa M, et al. Effect of nicotine on type 2 deiodinase activity in cultured rat glial cells. Endocr J 1999;46:107.

131. Canettieri G, Celi FS, Baccheschi G, et al. Isolation of human type 2 deiodinase gene promoter and characterization of a functional cyclic adenosine monophosphate response element. Endocrinology 2000;141:1804.

132. Borges M, Ingbar SH, Silva JE. Iodothyronine deiodinase activities in FRTL5 cells: predominance of type I 5′-deiodinase. Endocrinology 1990;126:3059.

133. Gereben B, Kollar A, Harney JW, et al. The mRNA structure has potent regulatory effects on type 2 iodothyronine deiodinase expression. Mol Endocrinol 2002;16:1667.

134. Burmeister LA, Pachucki J, St. Germain DL. Thyroid hormones inhibit type 2 iodothyronine deiodinase in the rat cerebral cortex by both pre- and posttranslational mechanisms. Endocrinology 1997;138:5231.

135. Leonard JL, Kaplan MM, Visser TJ, et al. Cerebral cortex responds rapidly to thyroid hormones. Science 1981;214:571.

136. Koenig RJ, Leonard JL, Senator D, et al. Regulation of thyroxine 5′-deiodinase activity by 3,5,3′-triiodothyronine in cultured rat anterior pituitary cells. Endocrinology 1984;115:324.

137. Silva JE, Leonard JL. Regulation of rat cerebrocortical and adenohypophyseal type II 5′-deiodinase by thyroxine, triiodothyronine, and reverse triiodothyronine. Endocrinology 1985;116: 1627.

138. Halperin Y, Shapiro LE, Surks MI. Down-regulation of type II L-thyroxine, 5′-monodeiodinase in cultured GC cells: different pathways of regulation by L-triiodothyronine and 3,3′,5′-triiodo-L-thyronine. Endocrinology 1994;135:1464.

139. Leonard JL, Silva JE, Kaplan MM, et al. Acute posttranscriptional regulation of cerebrocortical and pituitary iodothyronine 5′-deiodinases by thyroid hormone. Endocrinology 1984;114: 998.

140. Obregon MJ, Larsen PR, Silva JE. The role of 3,3′,5′-triiodothyronine in the regulation of type II iodothyronine 5′-deiodinase in the rat cerebral cortex. Endocrinology 1986;119: 2186.

141. St. Germain DL. The effects and interactions of substrates, inhibitors, and the cellular thiol-disulfide balance on the regulation of type II iodothyronine 5′-deiodinase. Endocrinology 1988;122:1860–1868.

142. Leonard JL, Siegrist-Kaiser CA, et al. Regulation of type II iodothyronine 5′-deiodinase by thyroid hormone: inhibition of actin polymerization blocks enzyme inactivation in cAMP-stimulated glial cells. J Biol Chem 1990;265:940.

143. Coux O, Tanaka K, Goldberg AL. Structure and functions of the 20S and 26S proteasomes. Annu Rev Biochem 1996;65:801.

144. Hershko A, Ciechanover A. The ubiquitin system. Annu Rev Biochem 1998;67:425.

145. Botero D, Gereben B, Goncalves C, et al. Ubc6p and Ubc7p are required for normal and substrate-induced endoplasmic reticulum-associated degradation of the human selenoprotein type 2 iodothyronine monodeiodinase. Mol Endocrinol 2002; 16:1999.

146. Tiwari S, Weissman AM. Endoplasmic reticulum (ER)-associated degradation of T cell receptor subunits: involvement of ER-associated ubiquitin-conjugating enzymes (E2s). J Biol Chem 2001;276:16193.

147. Kim BW, Zavacki AM, Curcio-Morelli C, et al. ER-associated degradation of the human type 2 iodothyronine deiodinase (D2) is mediated via an association between mammalian UBC7 and the carboxyl region of D2Mol Endocrinol 2003;17:2603.

148. Bianco AC, Harney J, Larsen PR, Identification of ubiquitinated forms of human type 2 deiodinase (hD2). 72nd Annual Meeting, American Thyroid Association, Palm Beach, FL, 1999, p 5.

149. Curcio-Morelli C, Zavacki AM, Christofollete M, et al. Deubiquitination of type 2 iodothyronine deiodinase by pVHL- interacting deubiquitinating enzymes regulates thyroid hormone activation. J Clin Invest 2003;112:189.

150. Bianco AC, Silva JE. Cold exposure rapidly induces virtual saturation of brown adipose tissue nuclear T3 receptors. Am J Physiol 1988;255:E496.

151. de Jesus LA, Carvalho SD, Ribeiro MO, et al. The type 2 iodothyronine deiodinase is essential for adaptive thermogenesis in brown adipose tissue. J Clin Invest 2001;108:1379.

152. Bianco AC, Silva JE. Intracellular conversion of thyroxine to triiodothyronine is required for the optimal thermogenic function of brown adipose tissue. J Clin Invest 1987;79:295.

153. Christoffolete MA, Linardi CCG, de Jesus LA, et al. Mice with targeted disruption of the Dio2 gene have cold-induced overexpression of uncoupling protein 1 gene but fail to increase brown adiose tissue lipogenesis and adaptive thermogenesis. Diabetes 2004;53:577.

154. Huang Y, Baker RT, Fischer-Vize JA. Control of cell fate by a deubiquitinating enzyme encoded by the fat facets gene. Science 1995;270:1828.

155. Moazed D, Johnson D. A deubiquitinating enzyme interacts with SIR4 and regulates silencing in S. cerevisiae. Cell 1996;86: 667.

156. Zhu Y, Carroll M, Papa FR, et al. DUB-1, a deubiquitinating enzyme with growth-suppressing activity. Proc Natl Acad Sci U S A 1996;93:3275.

157. Naviglio S, Mattecucci C, Matoskova B, et al. UBPY: a growth-regulated human ubiquitin isopeptidase. EMBO J 1998;17: 3241.

158. Park KC, Kim JH, Choi EJ, et al. Antagonistic regulation of myogenesis by two deubiquitinating enzymes, UBP45 and UBP69. Proc Natl Acad Sci U S A 2002;99:9733.

159. Li Z, Na X, Wang D, et al. Ubiquitination of a novel deubiquitinating enzyme requires direct binding to von Hippel-Lindau tumor suppressor protein. J Biol Chem 2002;277:4656.

160. Moreno M, Berry MJ, Horst C, et al. Activation and inactivation of thyroid hormone by type I iodothyronine deiodinase. FEBS Lett 1994;344:143.

161. Huang H, Marsh-Armstrong N, Brown DD. Metamorphosis is inhibited in transgenic Xenopus laevis tadpoles that overexpress type III deiodinase. Proc Natl Acad Sci U S A 1999;96:962.

162. Huang SA, Tu HM, Harney JW, et al. Severe hypothyroidism caused by type 3 iodothyronine deiodinase in infantile hemangiomas. N Engl J Med 2000;343:185.

163. Sato K, Robbins J. Thyroid hormone metabolism in cultured monkey hepatocarcinoma cells. J Biol Chem 1980;255:7347.

164. Kaplan MM, Yaskoski KA. Phenolic and tyrosyl ring deiodination of iodothyronines in rat brain homogenates. J Clin Invest 1980;66:551.

165. Kaplan MM, Yaskoski KA. Maturational patterns of iodothyronine phenolic tyrosyl ring deiodinase activities in rat cerebrum, cerebellem and hypothalamus. J Clin Invest 1981;67:1208.

166. Sorimachi K, Robbins J. Metabolism of thyroid hormones by cultured monkey hepatocarcinoma cells: nonphenolic ring deiodination and sulfation. J Biol Chem 1977;252:4458.

167. Richard K, Hume R, Kaptein E, et al. Ontogeny of iodothyronine deiodinases in human liver. J Clin Endocrinol Metab 1998; 83:2868.

168. Berry DL, Rose CS, Remo BF, et al. The expression pattern of thyroid hormone response genes in remodeling tadpole tissues defines distinct growth and resorption gene expression programs. Dev Biol 1998;203:24.

169. Hernandez A, Park JP, Lyon GJ, et al. Localization of the type 3 iodothyronine deiodinase (DIO3) gene to human chromosome 14q32 and mouse chromosome 12F1. Genomics 1998;53:119.

170. Hernandez A, Lyon GJ, Schneider MJ, et al. Isolation and characterization of the mouse gene for the type 3 iodothyronine deiodinase. Endocrinology 1999;140:124.

171. Hernandez A, Martinez ME, Croteau W, et al. Complex organization and structure of sense and antisense transcripts expressed from the DIO3 gene imprinted locus. Genomics 2004; 83:413.

172. Hernandez A, Fiering S, Martinez E, et al. The gene locus encoding iodothyronine deiodinase type 3 (Dio3) is imprinted in the fetus and expresses antisense transcripts. Endocrinology 2002;143:4483.

173. Salvatore D, Low SC, Berry M, et al. Type 3 iodothyronine deiodinase: cloning, in vitro expression, and functional analysis of the placental selenoenzyme. J Clin Invest 1995;96:2421.

174. Tu HM, Legradi G, Bartha T, et al. Regional expression of the type 3 iodothyronine deiodinase messenger ribonucleic acid in the rat central nervous system and its regulation by thyroid hormone. Endocrinology 1999;140:784.

175. Bates JM, Spate VL, Morris JS, et al. Effects of selenium deficiency on tissue selenium content, deiodinase activity, and thyroid hormone economy in the rat during development. Endocrinology 2000;141:2490.

176. Roti E, Braverman LE, Fang S-L, et al. Ontogenesis of placental inner ring thyroxine deiodinase and amniotic fluid 3,3′5′-triidothyronine concentration in the rat. Endocrinology 1982; 111:959.

177. Kaplan MM, McCann UD, Yaskoski KA, et al. Anatomical distribution of phenolic and tyrosyl ring iodothyronine deiodinases in the nervous system of normal and hypothyroid rats. Endocrinology 1981;109:397.

178. Galton VA, McCarthy PT, St Germain DL. The ontogeny of iodothyronine deiodinase systems in liver and intestine of the rat. Endocrinology 1991;128:1717.

179. Squire LR. Mechanisms of memory. Science 1986;342:1612.

180. Puymirat J, Miehe M, Marchand R, et al. Immunocytochemical localization of thyroid hormone receptors in the adult rat brain. Thyroid 1991;1:173.

181. Puymirat J. Thyroid receptors in the rat brain. Prog Neurobiol 1992;39:281.

182. Escamez MJ, Guadano-Ferraz A, Cuadrado A, et al. Type 3 iodothyronine deiodinase is selectively expressed in areas related to sexual differentiation in the newborn rat brain. Endocrinology 1999;140:5443.

183. Leonard JL, Larsen PR. Thyroid hormone metabolism in primary cultures of fetal rat brain cells. Brain Res 1985;327:1.

184. Cavalieri RR, Gavin LA, Cole R, et al. Thyroid hormone deiodinases in purified primary glial cell cultures. Brain Res 1986; 364:382.

185. Mori K, Yoshida K, Kayama T, et al. Thyroxine 5-deiodinase in human brain tumors. J Clin Endocrinol Metab 1993;77:1198.

186. Huang SA, Fish SA, Dorfman DM, et al. A 21-year-old woman with consumptive hypothyroidism due to a vascular tumor expressing type 3 iodothyronine deiodinase. J Clin Endocrinol Metab 2002;87:4457.

187. McCann UD, Shaw EA, Kaplan MM. Iodothyronine deiodination reaction types in several rat tissues: effects of age, thyroid status, and glucocorticoid treatment. Endocrinology 1984; 114:1513.

188. Marsh-Armstrong N, Huang H, Remo BF, et al. Asymmetric growth and development of the Xenopus laevis retina during metamorphosis is controlled by type III deiodinase. Neuron 1999;24:871.

189. Huang TS, Chopra IJ, Beredo A, et al. Skin is an active site for the inner ring monodeiodination of thyroxine to 3,3′,5′-triiodothyronine. Endocrinology 1985;117:2106.

190. Santini F, Vitti P, Chiovato L, et al. Role for inner ring deiodination preventing transcutaneous passage of thyroxine. J Clin Endocrinol Metab 2003;88:2825.

191. Castro MI, Braverman LE, Alex S, et al. Inner-ring deiodination of 3,5,3′-triiodothyronine in the in situ perfused guinea pig placenta. J Clin Invest 1985;76:1921.

192. Roti E, Fang SL, Green K, et al. Human placenta is an active site of thyroxine and 3,3′,5-triiodothyronine tyrosyl ring deiodination. J Clin Endocrinol Metab 1981;53:498.

193. Fay M, Roti E, Fang SL, et al. The effects of propylthiouracil, iodothyronines, and other agents on thyroid hormone metabolism in human placenta. J Clin Endocrinol Metab 1984;58:280.

194. Hidal JT, Kaplan MM. Characteristics of thyroxine 5′-deiodination in cultured human placental cells: regulation by iodothyronines. J Clin Invest 1985;76:947.

195. Galton VA, Martinez E, Hernandez A, et al. Pregnant rat uterus expresses high levels of the type 3 iodothyronine deiodinase. J Clin Audiometry 1999;103:979.

196. Wasco EC, Martinez E, Grant KS, et al. Determinants of iodothyronine deiodinase activities in rodent uterus. Endocrinology 2000;144:4253.

197. Huang SA, Dorfman DM, Genest DR, et al. Type 3 iodothyronine deiodinase is highly expressed in the human uteroplacental unit and in fetal epithelium. J Clin Endocrinol Metab 2003; 88:1384.

 

198. Esfandiari A, Courtin F, Lennon AM, et al. Induction of type III deiodinase activity in astroglial cells by thyroid hormones. Endocrinology 1992;131:1682.

199. Wassen FW, Schiel AE, Kuiper GG, et al. Induction of thyroid hormone-degrading deiodinase in cardiac hypertrophy and failure. Endocrinology 2002;143:2812.

200. Courtin F, Liva P, Gavaret JM, et al. Induction of 5-deiodinase activity in astroglial cells by 12-O-tetradecanoylphorbol 13-acetate and fibroblast growth factors. J Neurochem 1991;56:1107.

201. Pallud S, Ramauge M, Gavaret JM, et al. Regulation of type 3 iodothyronine deiodinase expression in cultured rat astrocytes: role of the Erk cascade. Endocrinology 1999;40:2917.

202. Klebanoff SJ, Green WL. Degradation of thyroid hormones by phagocytosing human leukocytes. J Clin Invest 1973;52:60.

203. Visvanathan A, Shanmugasundaram KR. Alterations in L-triiodothyronine aminotransferase activity in hypothyroid rats—effects of administration of iodobenzene and L-thyroxine. Ind J Exp Biol 1984;22:442.

204. Pittman CS, Shimizu T, Burger A, et al. The nondeiodinative pathways of thyroxine metabolism: 3,5,3′,5-tetraiodothyro acetic acid turnover in normal and fasting human subjects. J Clin Endocrinol Metab 1980;50:712.

205. Visser TJ. Role of sulfation in thyroid hormone metabolism. Chem Biol Interact 1994;92:293.

206. Visser TJ. Importance of deiodination and conjugation in the hepatic metabolism of thyroid hormone. In: Greer MA, ed. The thyroid gland. New York: Raven, 1990:255.

207. Sekura RD, Sato K, Cahnmann HJ, et al. Sulfate transfer to thyroid hormones and their analogs by hepatic aryl sulfotransferases. Endocrinology 1981;108:454.

208. Anderson RJ, Babbitt LL, Liebentritt DK. Human liver triiodothyronine sulfotransferase: copurification with phenol sulfotransferases. Thyroid 1995;5:61.

209. Sato K, Robbins J. Thyroid hormone metabolism in primary cultured rat hepatocytes. Effects of glucose, glucagon, and insulin. J Clin Invest 1981;68:475.

210. de Herder WW, Bonthuis F, Rutgers M, et al. Effects of inhibition of type I iodothyronine deiodinase and phenol sulfotransferase on the biliary clearance of triiodothyronine in rats. Endocrinology 1998;122:153.

211. Sorimachi K, Robbins J. Effects of propylthiouracil and methylmercaptoimidazol on metabolism of thyroid hormones by cultured monkey hepatocarcinoma cells. Horm Metab Res 1979;11:39.

212. Young WF Jr. Human liver tyrosylsulfotransferase. Gastroenterology 1990;99:1072.

213. Rooda SJE, Kaptein E, Visser TJ. Serum triiodothyronine sulfate in man measured by radioimmunoassay. J Clin Endocrinol Metab 1989;69:552.

214. Rooda SJE, Kaptein E, Rutgers M, et al. Increased plasma 3,5,3′-triiodothyronine sulfate in rats with inhibited type I iodothyronine dediodinase activity, as measured by radioimmunoassay. Endocrinology 1989;124:740.

215. van Stralen PG, van der Hoek HJ, Docter R, et al. Reduced T3 deiodination by the human hepatoblastoma cell line HepG2 caused by deficient T3sulfation. Biochim Biophys Acta 1993; 1157:114.

216. Chanoine JP, Safran M, Farwell AP, et al. Effects of selenium deficiency on thyroid hormone economy in rats. Endocrinology 1992;131:1787.

217. Wu SY, Huang WS, Polk D, et al. Identification of thyroxine sulfate (T4S) in human serum and amniotic fluid by a novel T4S radioimmunoassay. Thyroid 1992;2:101.

218. Chopra IJ, Santini F, Hurd RE, et al. A radioimmunoassay for measurement of thyroxine sulfate. J Clin Endocrinol Metab 1993;76:145.

219. Santini F, Cortelazzi D, Baggiani AM, et al. A study of the serum 3,5,3′-triiodothyronine sulfate concentration in normal and hypothyroid fetuses at various gestational stages. J Clin Endocrinol Metab 1993;76:1583.

220. Wu S-Y, Huang W-S, Polk D, et al. The development of a radioimmunoassay for reverse triiodothyronine sulfate in human serum and amniotic fluid. J Clin Endocrinol Metab 1993;76: 1625.

221. LoPresti JS, Nicoloff JT. 3,5,3′-Triiodothyronine (T3) sulfate: a major metabolite in T3 metabolism in man. J Clin Endocrinol Metab 1994;78:688.

222. Frumess RD, Larsen PR. Correlation of serum triiodothyronine (T3) and thyroxine (T4) with biologic effects of thyroid hormone replacement in propylthiouracil-treated rats. Metabolism 1975;24:547.

223. Larsen PR, Frumess RD. Comparison of the biological effects of thyroxine and triiodothyronine in the rat. Endocrinology 1977;100:980.

224. Silva JE, Larsen PR. Peripheral metabolism of homologous thyrotropin in euthyroid and hypothyroid rats: acute effects of thyrotropin-releasing hormone, triiodothyronine, and thyroxine. Endocrinology 1978;102:1783.

225. Larsen PR, Dick TE, Markovitz BP, et al. Inhibition of intrapituitary thyroxine to 3,5,3′-triiodothyronine conversion prevents the acute suppression of thyrotropin release by thyroxine in hypothyroid rats. J Clin Invest 1979;64:117.

226. Riesco G, Taurog A, Larsen R, et al. Acute and chronic responses to iodine deficiency in rats. Endocrinology 1977;100: 303.

227. Segerson TP, Kauer J, Wolfe H, et al. Thyroid hormone regulates TRH biosynthesis in the paraventricular nucleus of the rat hypothalamus. Science 1987;238:78.

228. Connors JM, Hedge GA. Feedback effectiveness of periodic versus constant triiodothyronine replacement. Endocrinology 1980;106:911.

229. Connors JM, Hedge GA. Feedback regulation of thyrotropin by thyroxine under physiological conditions. Am J Physiol 1981;240:E308.

230. Kakucska I, Rand W, Lechan RM. Thyrotropin-releasing hormone (TRH) gene expression in the hypothalamic paraventricular nucleus is dependent upon feedback regulation by both triiodothyronine and thyroxine. Endocrinology 1992;130:2845.

231. Izumi M, Larsen PR. Triiodothyronine, thyroxine, and iodine in purified thyroglobulin from patients with Graves' disease. J Clin Invest 1977;59:1105.

232. Larsen PR. Thyroidal triiodothyronine and thyroxine in Graves' disease: correlation with presurgical treatment, thyroid status, and iodine content. J Clin Endocrinol Metab 1975;41: 1098.

233. Laurberg P. Mechanisms governing the relative proportions of thyroxine and 3,5,3′-triiodothyronine in thyroid secretion. Metabolism 1984;33:379.

234. Geffner DL, Azukizawa M, Hershman JM. Propylthiouracil blocks extrathyroidal conversion of thyroxine to triiodothyronine and augments thyrotropin secretion in man. J Clin Invest 1975;55:224.

235. Saberi M, Sterling FH, Utiger RD. Reduction in extrathyroidal triiodothyronine production by propylthiouracil in man. J Clin Invest 1975;55:218.

236. LoPresti JS, Eigen A, Kaptein E, et al. Alterations in 3,3′5′-triiodothyronine metabolism in response to propylthiouracil, dexamethasone, and thyroxine administration in man. J Clin Invest 1989;84:1650.

237. Inada M, Kasagi K, Kurata S, et al. Estimation of thyroxine and triiodothyronine distribution and of the conversion rate of thyroxine to triiodothyronine in man. J Clin Invest 1975;55:1337.

238. Lum SM, Nicoloff JT, Spencer CA, et al. Peripheral tissue mechanism for maintenance of serum triiodothyronine values in a thyroxine-deficient state in man. J Clin Invest 1984;73: 570.

239. Nicoloff JT, Lum SM, Spencer CA, et al. Peripheral autoregulation of thyroxine to triiodothyronine conversion in man. Horm Metab Res 1984;14:74(suppl.).

240. Pilo A, Iervasi G, Vitek F, et al. Thyroidal and peripheral production of 3,5,3′-triiodothyronine in humans by multicompartmental analysis. Am J Physiol 1990;258:E715.

241. Oppenheimer JH, Schwartz HL, Surks MI. Propylthiouracil inhibits the conversion of L-thyroxine to L-triiodothyronine: an explanation of the antithyroxine effect of propylthiouracil and evidence supporting the concept that triiodothyronine is the active thyroid hormone. J Clin Invest 1972;51:2493.

242. Nguyen TT, Chapa F, DiStefano JJ 3rd. Direct measurement of the contributions of type I and type II 5′-deiodinases to whole body steady state 3,5,3′-triiodothyronine production from thyroxine in the rat. Endocrinology 1998;139:4626.

243. Hosoi Y, Murakami M, Mizuma H, et al. Expression and regulation of type II iodothyronine deiodinase in cultured human skeletal muscle cells. J Clin Endocrinol Metab 1999;84:3293.

244. Friesema EC, Docter R, Moerings EP, et al. Identification of thyroid hormone transporters. Biochem Biophys Res Commun 1999;254:497.

245. Hennemann G, Docter R, Friesema ECH, et al. Plasma membrane transport of thyroid hormones and its role in thyroid hormone metabolism and bioavailability. Endocr Rev 2001;22: 451.

246. Oppenheimer JH. Thyroid hormone action at the cellular level. Science 1979;203:971.

247. Silva JE, Leonard JL, Crantz FR, et al. Evidence for two tissue specific pathways for in vivo thyroxine 5′ deiodination in the rat. J Clin Invest 1982;69:1176.

248. Bianco AC, Silva JE. Nuclear 3,5,3′-triiiodothyronine (T3) in brown adipose tissue: receptor occupancy and sources of T3 as determined by in vivo techniques. Endocrinology 1897;120:55.

249. van Doorn JD, Roelfsema F, van der Heide D. Contribution from local conversion of thyroxine to 3,5,3′-triiodothyronine to cellular 3,5,3′-triiodothyronine in several organs in hypothyroid rats at isotope equilibrium. Acta Endocrinol (Copenh) 1982;101:386.

250. van Doorn JD, van der Heide D, Roelfsema F. Sources and quantity of 3,5,3′-triiodothyronine in several tissues of the rat. J Clin Invest 1983;72:1778.

251. van Doorn JD, Roelfsema F, van der Heide D. Concentrations of thyroxine and 3,5,3′-triiodothyronine at 34 different sites in euthyroid rats as determined by an isotopic equilibrium technique. Endocrinology 1985;117:1201.

252. Eales JG, McLeese JM, Holmes JA, et al. Changes in intestinal and hepatic thyroid hormone deiodination during spontaneous metamorphosis of the sea lamprey, Petromyzon marinus. J Exp Zool 2000;286:305.

253. Larsen PR, Bavli SZ, Castonguay M, et al. Direct radioimmunoassay of nuclear 3,5,3′ triiodothyronine in rat anterior pituitary. J Clin Invest 1980;65:675.

254. Oppenheimer JH, Schwartz HL. Stereospecific transport to triiodothyronine from plasma to cytosol and from cytosol to nucleus in rat liver, kidney, brain and heart. J Clin Invest 1985; 75:147.

255. Dunn JT. Global IDD status. IDD Newsletter 1999;15:17.

256. Silva JE. Disposal rates of thyroxine and triiodothyronine in iodine-deficient rats. Endocrinology 1972;91:1430.

257. Pazos-Moura CC, Moura EG, Dorris ML, et al. Effect of iodine deficiency and cold exposure on thyroxine 5′-deiodinase activity in various rat tissues. Am J Physiol 1991;260:E175.

258. Okamura K, Taurog A, Krulich L. Hypothyroidism in severely iodine-deficient rats. Endocrinology 1981;109:464.

259. Santisteban P, Obregon MJ, Rodriguez-Pena A, et al. Are iodine-deficient rats euthyroid? Endocrinology 1982;110:1780.

260. Obregon MJ, Santisteban P, Rodriguez-Pena A, et al. Cerebral hypothyroidism in rats with adult-onset iodine deficiency. Endocrinology 1984;115:614.

261. Riesco G, Taurog A, Larsen PR. Variations in the response of the thyroid gland of the rat to different low-iodine diets: correlation with iodine content of diet. Endocrinology 1976; 99:270.

262. Greer MA, Grimm Y, Studer H. Qualitative changes in the secretion of thyroid hormones induced by iodine deficiency. Endocrinology 1968;83:1193.

263. Abrams GM, Larsen PR. Triiodothyronine and thyroxine in the serum and thyroid glands of iodine-deficient rats. J Clin Invest 1873;52:2522.

264. Silva JE, Gordon MB, Crantz FR, et al. Qualitative and quantitative differences in the pathways of extrathyroidal triiodothyronine generation between euthyroid and hypothyroid rats. J Clin Invest 1984;73:898.

265. Campos-Barros A, Meinhold H, Walzog B, et al. Effects of selenium and iodine deficiency on thyroid hormone concentrations in the central nervous system of the rat. Eur J Endocrinol 1997;136:316.

266. Guadano-Ferraz A, Escamez MJ, Rausell E, et al. Expression of type 2 iodothyronine deiodinase in hypothyroid rat brain indicates an important role of thyroid hormone in the development of specific primary sensory systems. J Neurosci 1999; 19:3430.

267. Peeters R, Fekete C, Goncalves C, et al. Regional physiological adaptation of the central nervous system deiodinases to iodine deficiency. Am J Physiol Endocrinol Metab 2001;281: E54.

268. Kim SW, Harney JW, Larsen PR. Studies of the hormonal regulation of type 2 5′-iodothyronine deiodinase messenger ribonucleic acid in pituitary tumor cells using semiquantitative reverse transcription-polymerase chain reaction. Endocrinology 1998;139:4895.

269. St. Germain DL, Schwartzman RA, Croteau W, et al. A thyroid hormone-regulated gene in Xenopus laevis encodes a type III iodothyronine 5-deiodinase. Proc Natl Acad U S A 1994;91: 7767.

270. Schroder-van der Elst JP, van der Heide D, Morreale de Escobar G, et al. Iodothyronine deiodinase activities in fetal rat tissues at several levels of iodine deficiency: a role for the skin in 3,5,3′-triiodothyronine economy? Endocrinology 1998;139: 2229.

271. Silva JE, Matthews PS. Production rates and turnover of triiodothyronine in rat developing cerebral cortex and cerebellum. J Clin Invest 1984;74:1035.

272. Crantz FR, Silva JE, Larsen PR. An analysis of the sources and quantity of 3,5,3′-triiodothyronine specifically bound to nuclear receptors in rat cerebral cortex and cerebellum. Endocrinology 1982;110:367.

273. Silva JE, Larsen PR. Comparison of iodothyronine 5′-deiodinase and other thyroid-hormone-dependent enzyme activities in the cerebral cortex of hypothyroid neonatal rat: evidence for adaptation to hypothyroidism. J Clin Invest 1982;70:1110.

274. Porterfield SP, Hendrich CE. The role of thyroid hormones in prenatal and neonatal neurological development—current perspectives. Endocr Rev 1993;14:94.

275. Pasquini JM, Adamo AM. Thyroid hormones and the central nervous system. Dev Neurosci 1994;16:1.

276. Brent GA. The molecular basis of thyroid hormone action. N Engl J Med 1994;331:847.

 

277. Zhang J, Lazar MA. The mechanism of action of thyroid hormones. Annu Rev Physiol 2000;62:439.

278. Kaplan MM, Shaw EA. Type II iodothyronine 5′-deiodination by human and rat placenta in vitro. J Clin Endocrinol Metab 1984;59:253.

279. Kaplan MM. Regulatory influences on iodothyronine deiodination in animal tissues. In: Hennemann G, ed. Thyroid hormone metabolism. New York: Marcel Dekker, 1986:231.

280. Marsh-Armstrong N, Cai L, Brown DD. Thyroid hormone controls the development of connections between the spinal cord and limbs during Xenopus laevis metamorphosis. Proc Natl Acad Sci U S A 2004;101:165.

281. Cai L, Brown DD. Expression of type II iodothyronine deiodinase marks the time that a tissue responds to thyroid hormone-induced metamorphosis inXenopus laevis. Dev Biol 2004;266: 87.

282. Huang T, Chopra IJ, Boado R, et al. Thyroxine inner ring monodeiodinating activity in fetal tissues of the rat. Pediatr Res 1988;23:196.

283. Ng L, Goodyear RJ, Woods CA, et al. Hearing loss and retarded cochlear development in mice lacking type 2 iodothyronine deiodinase. Proc Natl Acad Sci U S A 2004;101:3474.

284. Thorpe-Beeston JG, Nicolaides KH, McGregor AM. Fetal thyroid function. Thyroid 1992;2:207.

285. Contempre B, Jauniaux E, Calvo R, et al. Detection of thyroid hormones in human embryonic cavities during the first trimester of pregnancy. J Clin Endocrinol Metab 1993;77:1719.

286. Fisher DA, Lehman H, Lackey C. Placental transport of thyroxine. J Clin Endocrinol 1964;24:393.

287. Abuid J, Klein AH, Foley TP Jr, et al. 1974 Total and free triiodothyronine and thyroxine in early infancy. J Clin Endocrinol Metab 1974;39:263.

288. Chopra IJ, Wu SY, Chua Teco GN, et al. A radioimmunoassay for measurement of 3,5,3′-triiodothyronine sulfate: studies in thyroidal and non-thyroidal disease, pregnancy and neonatal life. J Clin Endocrinol Metab 1992;75:189.

289. Santini F, Chopra IJ, Wu SY, et al. Metabolism of 3,5,3′-triiodothyronine sulfate by tissues of the fetal rat: a consideration of the role of desulfation of 3,5,3′-triiodothyronine sulfate as a source of T3Pediatr Res 1992;31:541.

290. Koopdonk-Kool JM, deVijlder JJM, Veenboer GJM, et al. Type II and Type III deiodinase activity in human placenta as a function of gestational age. J Clin Endocrinol Metab 1996;81: 2154.

291. Stulp MR, de Vijlder JJ, Ris-Stalpers C. Placental iodothyronine deiodinase III and II ratios, mRNA expression compared to enzyme activity. Mol Cell Endocrinol 1998;142:67.

292. Yoshida K, Suzuki M, Sakurada T. Changes in thyroxine monodeiodination in rat liver, kidney and placenta during pregnancy. Acta Endocrinol (Copenh) 1984;107:495.

293. Remesar X, Arola L, Palou A, et al. Activities of enzymes involved in amino-acid metabolism in developing rat placenta. Eur J Biochem 1980;110:289.

294. Mortimer RH, Galligan JP, Cannell GR, et al. Maternal to fetal thyroxine transmission in the human term placenta is limited by inner ring deiodination. J Clin Endocrinol Metab 1996;81: 2247.

295. Bradley DJ, Towle HC, Young WSI. Spatial and termporal expression of a- and b-thyroid hormone receptor mRNAs, including the b2-subtype, in the developing mammalian nervous system. J Neurosci 1992;12:2288.

296. Kraft JC, Willhite CC, Juchau MR. Embryogenesis in cultured whole rat embryos after combined exposures to 3,3′,5-triiodo L-thyronine (T3) plus all-trans-retinoic acid and to T3 plus 9-cis-retinoic acid. J Craniofac Genet Dev Biol 1994; 14:75.

297. Klein AH, Hobel CJ, Sack J, et al. Effect of intraamniotic fluid thyroxine injection on fetal serum and amniotic fluid iodothyronine concentrations. J Clin Endocrinol Metab 1978; 47:1034.

298. Mandel SJ, Larsen PR, Seely EW, et al. Increased need for thyroxine during pregnancy in women with primary hypothyroidism. N Engl J Med 1990;323:91.

299. Vulsma T, Gons MH, DeVijlder JMM. Maternal fetal transfer of thyroxine in congenital hypothyroidism due to a total organification defect of thyroid dysgenesis. N Engl J Med 1989;321: 13.

300. LaFranchi S. Congenital hypothyroidism: etiologies, diagnosis, and management. Thyroid 1999;9:735.

301. Ayling RM, Davenport M, Hadzic N, et al. Hepatic hemangioendothelioma associated with production of humoral thyrotropin-like factor. J Pediatr 2001;138:932.

302. Kaplan MM. Monitoring thyroxine treatment during pregnancy. Thyroid 1992;2:147.

303. Alexander EK, Marqusee E, Lawrence J, et al. Timing and magnitude of increases in levothyroxine requirements during pregnancy in women with hypothyroidism. N Engl J Med 2004; 241.

304. Ishii H, Inada M, Tanaka K, et al. Triiodothyronine generation from thyroxine in human thyroid: enhanced conversion in Graves' thyroid tissue. J Clin Endocrinol Metab 1981;52: 1211.

305. Sugawara M, Lau R, Wasser HL, et al. Thyroid T45′-deiodinase activity in normal and abnormal human thyroid glands. Metabolism 1984;33:332.

306. Wu SY, Shyh TP, Chopra IJ, et al. Comparison sodium ipodate (oragrafin) and propylthiouracil in early treatment of hyperthyroidism. J Clin Endocrinol Metab 1982;54:630.

307. Burgi H, Wimpfheimer C, Burger A, et al. Changes of circulating thyroxine, triiodothyronine and reverse triiodothyronine after radiographic contrast agents. J Clin Endocrinol Metab 1976;43:1203.

308. Croxson MS, Hall TD, Nicoloff JT. Combination drug therapy for treatment of hyperthyroid Graves' disease. J Clin Endocrinol Metab 1977;45:623.

309. Laurberg P, Torring J, Weeke J. A comparison of the effects of propylthiouracil and methimazol on circulating thyroid hormones and various measures of peripheral thyroid hormone effects in thyrotoxic patients. Acta Endorinol (Copenh) 1985; 108:51.

310. Kim BW, Daniels GH, Harrison BJ, et al. Overexpression of type 2 iodothyronine deiodinase in follicular carcinoma as a cause of low circulating free thyroxine levels. J Clin Endocrinol Metab 2003;88:594.