SNAPSHOT

Key Event Component

Domain of Applicability

Key Event Component

Domain of Applicability

Key Event Description

Thyroid hormones (TH) are present in brain tissue of most vertebrate species, and thyroxine (T4) is converted to triiodothyronine locally in this tissue.  The amount of THs in brain is known to vary during development and to differ among brain regions (Calvo et al., 1990; Kester et al., 2004; Tu et al., 1999). In human cerebral cortex, T3 increases steadily from 13-weeks, reaching adult levels by 20 weeks post conception. This occurs despite very low and unchanging levels in fetal serum T3, when fetal serum T4 increases 3-fold over the same period. This indicates that T3 in fetal brain is locally generated from serum-derived T4 via the activity of deiodinases, primarily DIO2. DIO2 serves to convert T4 to T3. During this time in fetal development DIO3 activity, which converts T3 to the inactive reverse T3 (rT3), remains very low in cortex.  In contrast, in other brain regions including hippocampus and cerebellum, T3 remains low throughout early and mid-gestation and corresponds with high activity of DIO3 in these brain regions. In late gestation and after birth, DIO3 levels drop in hippocampus and cerebellum with a corresponding increase in T3 concentrations (Kester et al., 2004). 

A similar spatial and temporal profile of deiodinase activity and corresponding brain hormone concentrations has been observed in rodent brain (Calvo et al., 1990; Tu et al., 1999). In the rat, either whole brain or cortex have been preferentially assessed due to the low levels of hormones present and the small tissue volumes make quantitification difficult. Brain T3 and T4 rise in parallel from gestational day 10 to gestational day 20 in rat. They are typically both quite low until gestational 17 with steep increases between GD18 and GD20 corresponding to the onset of fetal thyroid function (Calvo et al., 1990; Ruiz de Ono et al., 1988; Obergon et al., 1981). Just before birth, brain T3 and T4 concentrations are about one-third to one-half that of adult brain. Brain development in the early postnatal period in rat is roughly equivalent to the 3^rd trimester in humans such that adult levels of T3 and T4 in brain are not reached in rodents until the 2^nd-3^rd postnatal week.

For THs to gain access to brain tissue they need to cross the blood brain barrier (BBB) which regulates the active transport of TH into neurons. Many transporter proteins have been identified, and the monocarboxylate transporters (Mct8, Mct10) and anion-transporting polypeptide (OATP1c1) show the highest degree of affinity towards TH and are prevalent in brain (Jansen et al., 2007; Mayer et al., 2014).  Transporters express a distinct distribution pattern that varies by tissue and age (Friesema et al., 2005; Henneman et al., 2001; Visser et al., 2007; Heuer et al., 2005; Muller and Heuer, 2007). Although several transporters have been identified, current knowledge of cell specific profile of transporters is limited. 

Most of the hormone transported across the blood brain barrier is in the form of T4, primarily though the cellular membrane transporters (e.g., OATP1c1 transporter) into the astrocyte (Visser and Visser, 2012; Sugiyama et al., 2003; Tohyama et al., 2004). Within the astrocyte, T4 is converted into T3 via the local activity of deiodinase 2 (DIO2) (Guadano-Ferraz et al., 1997).  A small amount of T3 may cross the blood brain barrier directly via the T3-specific transporter, MCT8 (Heuer et al., 2005). Although in mature brain T3 derives partially from the circulation and from the deiodination of T4, in the fetal brain T3 is exclusively a product of T4 deiodination (Calvo et al., 1990; Grijota-Martinez et al., 2011). In both cases, only the required amount of T3 is utilized in neurons and the excess is degraded by the neuron-specific deiodinase DIO3 (Tu et al., 1999; St. Germain et al., 2009; Hernandez et al., 2010).

Both deiodinase and transporter expression in brain peak in different brain regions at different times in fetal and neonatal life (Kester et al., 2004; Bates et al., 1999; Muller and Heuer, 2014; Heuer, 2007). Collectively, these spatial and temporal patterns of transporter expression and deiodinase activity provide exquisite control of brain T3 available for nuclear receptor activation and regulated gene expression.

How it is Measured or Detected

Radioimmunoassays (RIAs) are commonly used to detect TH in the brain (e.g., Obregon et al., 1982; Calvo et al., 1990; Morse et al., 1996; Bansal et al., 2005; Gilbert et al., 2013). The method (and minor variants) is well established in the published literature. However, it is not available in a simple 'kit' and requires technical knowledge of RIAs, thus has not been used in most routine toxicology studies. Evaluations in neuronal tissue are complicated by the difficulty of the fatty matrix, heterogeneity of regions within the brain, and low tissue concentrations and small tissue amounts especially in immature brain. Most often whole brain homogenates are assessed, obfuscating the known temporal and regional differences in brain hormone present. Two analytical techniques, LC- and HPLC-inductively coupled plasma–mass spectrometry have recently been used to measure brain concentrations of TH. These techniques have proven capable of measuring very low levels in whole-body homogenates of frog tadpoles at different developmental stages (e.g., Simon et al., 2002; Tietge et al., 2010). The assay detects I–, MIT, DIT, T4, T3, and rT3. More recently, Wang and Stapleton (2010) and Donzelli et al. (2016) used liquid chromatography-tandem mass spectrometry for the simultaneous analysis of five THs including thyroxine (T4), 3,3′,5-triidothyronine (T3), 3,3′,5′-triiodothyronine (rT3; reverse T3), 3,3′-diiodothyronine (3,3′-T2), and 3,5-diiodothyronine (3,5-T2) in serum and a variety of tissues including brain. These analytical methods require expensive equipment and technical expertise and as such are not routinely used.

References

Bansal R, You SH, Herzig CT, Zoeller RT (2005). Maternal thyroid hormone increases HES expression in the fetal rat brain: an effect mimicked by exposure to a mixture of polychlorinated biphenyls (PCBs). Brain Res Dev Brain Res 156:13-22.

Bates JM, St Germain DL, Galton VA. Expression profiles of the three iodothyronine deiodinases, D1, D2, and D3, in the developing rat. Endocrinology. 1999 Feb;140(2):844-51.

Bernal J. (2013). Thyroid Hormones in Brain Development and Function.  In: De Groot LJ, Chrousos G, Dungan K, Feingold KR, Grossman A, Hershman JM, Koch C, Korbonits M, McLachlan R, New M, Purnell J, Rebar R, Singer F, Vinik A, editors. Endotext [Internet]. South Dartmouth (MA): MDText.com, Inc.; 2000-2015. www.thyroidmanager.org

Calvo R, Obregón MJ, Ruiz de Oña C, Escobar del Rey F, Morreale de Escobar G. (1990). Congenital hypothyroidism, as studied in rats. Crucial role of maternal thyroxine but not of 3,5,3′-triiodothyronine in the protection of the fetal brain. J. Clin. Invest. 86:889-899.

Chatonnet F., Picou F., Fauquier T., and Flamant F., (2011). Thyroid Hormone Action in Cerebellum and Cerebral Cortex Development, Journal of Thyroid Research, Volume 2011, Article ID 145762, 8 pages http://dx.doi.org/10.4061/2011/145762)

Denver, RJ 1998 The molecular basis of thyroid hormone-dependent central nervous system remodeling during amphibian metamorphosis. Comparative Biochemistry and Physiology Part C: Pharmacology, Toxicology and Endocrinology, 119:219-228.

Donzelli R, Colligiani D, Kusmic C, Sabatini M, Lorenzini L, Accorroni A, Nannipieri M, Saba A, Iervasi G, Zucchi R. Effect of Hypothyroidism and Hyperthyroidism on Tissue Thyroid Hormone Concentrations in Rat. Eur Thyroid J. 2016 Mar;5(1):27-34.

Friesema EC, Jansen J, Milici C, Visser TJ (2005) Thyroid hormone transporters. Vitam Horm 70:137-167.

Galton VH 1983 Thyroid hormone action in amphibian metamorphosis. In: Oppenheimer JH, Samuels HH (eds) Molecular Basis of Thyroid Hormone Action. Academic Press, New York, pp 445–483.

Gilbert ME, Hedge JM, Valentin-Blasini L, Blount BC, Kannan K, Tietge J, Zoeller RT, Crofton KM, Jarrett JM, Fisher JW (2013) An animal model of marginal iodine deficiency during development: the thyroid axis and neurodevelopmental outcome. Toxicol Sci 132:177-195.

Grijota-Martinez C, Diez D, Morreale de Escobar G, Bernal J, Morte B. (2011). Lack of action of exogenously administered T3 on the fetal rat brain despite expression of the monocarboxylate transporter 8. Endocrinology. 152:1713-1721.

Guadano-Ferraz A, Obregon MJ, St Germain DL, Bernal J. (1997). The type 2 iodothyronine deiodinase is expressed primarily in glial cells in the neonatal rat brain. Proc Natl Acad Sci USA. 94: 10391–10396.

Hennemann G, Docter R, Friesema EC, de Jong M, Krenning EP, Visser TJ. (2001). Plasma membrane transport of thyroid hormones and its role in thyroid hormone metabolism and bioavailability. Endocr Rev. 22:451-476.

Hernandez A, Quignodon L, Martinez ME, Flamant F, St Germain DL. Type 3 deiodinase deficiency causes spatial and temporal alterations in brain T3 signaling that are dissociated from serum thyroid hormone levels. Endocrinology. 2010 Nov;151(11):5550-8.

Heuer H. (2007). The importance of thyroid hormone transporters for brain development and function. Best Pract Res Clin Endocrinol Metab. 21:265–276.

Heuer H, Maier MK, Iden S, Mittag J, Friesema EC, Visser TJ, Bauer K. (2005). The monocarboxylate transporter 8 linked to human psychomotor retardation is highly expressed in thyroid hormone-sensitive neuron populations. Endocrinology 146:1701–1706.

Jansen J, Friesema EC, Kester MH, Milici C, Reeser M, Gruters A, Barrett TG, Mancilla EE, Svensson J, Wemeau JL, Busi da Silva Canalli MH, Lundgren J, McEntagart ME, Hopper N, Arts WF, Visser TJ (2007) Functional analysis of monocarboxylate transporter 8 mutations identified in patients with X-linked psychomotor retardation and elevated serum triiodothyronine. J Clin Endocrinol Metab 92:2378-2381.

Kester MH, Martinez de Mena R, Obregon MJ, Marinkovic D, Howatson A, Visser TJ, Hume R, Morreale de Escobar G. (2004). Iodothyronine levels in the human developing brain: major regulatory roles of iodothyronine deiodinases in different areas. J Clin Endocrinol Metab 89:3117–3128.

Mayer S, Müller J, Bauer R, Richert S, Kassmann CM, Darras VM, Buder K, Boelen A, Visser TJ, Heuer H. Transporters MCT8 and OATP1C1 maintain murine brain thyroid hormone homeostasis. J Clin Invest. 2014 May 1;124(5):1987-99.

Moog N.K., Entringer S., Heim Ch., Wadhwa PD., Kathmann N., Buss C. (2017). Influence of maternal thyroid hormones during gestation on fetal  brain development. Neuroscience, 2017, 342: 68–100. doi:10.1016/j.neuroscience.2015.09.0

Morse DC, Wehler EK, Wesseling W, Koeman JH, Brouwer A. Alterations in rat brain thyroid hormone status following pre- and postnatal exposure to polychlorinated biphenyls (Aroclor 1254). Toxicol Appl Pharmacol. 1996 Feb;136(2):269-79

Müller J, Heuer H. Expression pattern of thyroid hormone transporters in the postnatal mouse brain. Front Endocrinol (Lausanne). 2014 Jun 18:5:92.

Obregon MJ, Mallol J, Escobar del Rey F, Morreale de Escobar G. (1981). Presence of l-thyroxine and 3,5,3-triiodo-l-thyronine in tissues from thyroidectomised rats. Endocrinology 109:908-913.

Power DM, Llewellyn L, Faustino M, Nowell MA, Björnsson BT, Einarsdottir IE, Canario AV, Sweeney GE. Thyroid hormones in growth and development of fish. Comp Biochem Physiol C Toxicol Pharmacol. 2001 Dec;130(4):447-59.

Ruiz de Oña C, Obregón MJ, Escobar del Rey F, Morreale de Escobar G. Developmental changes in rat brain 5'-deiodinase and thyroid hormones during the fetal period: the effects of fetal hypothyroidism and maternal thyroid hormones. Pediatr Res. 1988 Nov;24(5):588-94.

Simon R, Tietge JE, Michalke B, Degitz S, Schramm KW. Iodine species and the endocrine system: thyroid hormone levels in adult Danio rerio and developing Xenopus laevis. Anal Bioanal Chem. 2002 Feb;372(3):481-5.

St Germain DL, Galton VA, Hernandez A. (2009). Minireview: Defining the roles of the iodothyronine deiodinases: current concepts and challenges. Endocrinology. 150:1097-107.

Sugiyama D, Kusuhara H, Taniguchi H, Ishikawa S, Nozaki Y, Aburatani H, Sugiyama Y. (2003). Functional characterization of rat brain-specific organic anion transporter (Oatp14) at the blood–brain barrier: high affinity transporter for thyroxine. J Biol Chem. 278:43489–43495.

Tietge JE, Butterworth BC, Haselman JT, Holcombe GW, Hornung MW, Korte JJ, Kosian PA, Wolfe M, Degitz SJ. Early temporal effects of three thyroid hormone synthesis inhibitors in Xenopus laevis. Aquat Toxicol. 2010 Jun 1;98(1):44-50.

Tohyama K, Kusuhara H, Sugiyama Y. (2004). Involvement of multispecific organic anion transporter, Oatp14 (Slc21a14), in the transport of thyroxine across the blood-brain barrier. Endocrinology. 145: 4384–4391.

Tu HM, Legradi G, Bartha T, Salvatore D, Lechan RM, Larsen PR. (1999). Regional expression of the type 3 iodothyronine deiodinase messenger ribonucleic acid in the rat central nervous system and its regulation by thyroid hormone. Endocrinology. 140: 784–790.

Van Herck SL, Geysens S, Delbaere J, Darras VM. Regulators of thyroid hormone availability and action in embryonic chicken brain development. Gen Comp Endocrinol. 2013.190:96-104.

Visser EW, Visser TJ. (2012). Finding the way into the brain without MCT8. J Clin Enodcrinol Metab. 97:4362-4365.

Visser WE, Friesema EC, Jansen J, Visser TJ. (2007). Thyroid hormone transport by monocarboxylate transporters. Best Pract Res Clin Endocrinol Metab. 21:223–236.

Wang, D. and Stapleton, HM. (2010) Analysis of thyroid hormones in serum by liquid chromatography -tandem mass spectrometry. Anal Bioanal Chem. 2010 Jul; 397(5): 1831–1839

Key Event Component

Evidence for Perturbation by Stressor

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Propylthiouracil

6-n-propylthouracil is a classic positive control for inhibition of TPO

Perchlorate

Perchlorate ion (ClO− ₄) is a classic positive control for inhibition of NIS

Methimazole

Classic positive control

Domain of Applicability

Key Event Description

All iodothyronines are derived from the modification of tyrosine molecules (Taurog, 2000). There are two biologically active thyroid hormones (THs) in serum, triiodothyronine (T3) and T4, and a few inactive iodothyronines (rT3, 3,5-T2).  T4 is the predominant TH in circulation, comprising approximately 80% of the TH excreted from the thyroid gland and is the pool from which the majority of T3 in serum is generated (Zoeller et al., 2007). As such, serum T4 changes usually precede changes in other serum THs.  Decreased thyroxine (T4) in serum results result from one or more MIEs upstream and is considered a key biomarker of altered TH homeostasis (DeVito et al., 1999). 

Serum T4 is used as a biomarker of TH status because the circulatory system serves as the major transport and delivery system for TH delivery to tissues. The majority of THs in the blood are bound to transport proteins (Bartalena and Robbins, 1993). In serum, it is the unbound, or ‘free’ form of the hormone that is thought to be available for transport into tissues. Free hormones are approximately 0.03 and 0.3 percent for T4 and T3, respectively. There are major species differences in the predominant binding proteins and their affinities for THs (see below). However, there is broad agreement that changes in serum concentrations of THs is diagnostic of thyroid disease or chemical-induced disruption of thyroid homeostasis (DeVito et al., 1999; Miller et al., 2009; Zoeller et al., 2007).

Normal serum T4 reference ranges can be species and lifestage specific. In rodents, serum THs are low in the fetal circulation, increasing as the fetal thyroid gland becomes functional on gestational day 17, just a few days prior to birth. After birth serum hormones increase steadily, peaking at two weeks, and falling slightly to adult levels by postnatal day 21 (Walker et al., 1980; Harris et al., 1978; Goldey et al., 1995; Lau et al., 2003). Similarly, in humans, adult reference ranges for THs do not reflect the normal ranges for children at different developmental stages, with TH concentrations highest in infants, still increased in childhood, prior to a decline to adult levels coincident with pubertal development (Corcoran et al. 1977; Kapelari et al., 2008). In some frog species, there is an analogous peak in thyroid hormones in tadpoles that starts around NF stage 56, peaks at Stage 62 and the declines to lower levels by Stage 56 (Sternberg et al., 2011; Leloup and Buscaglia, 1977). 

How it is Measured or Detected

Serum T3 and T4 can be measured as free (unbound) or total (bound + unbound). Free hormone concentrations are clinically considered more direct indicators of T4 and T3 activities in the body, but in animal studies, total T3 and T4 are typically measured. Historically, the most widely used method in toxicology is radioimmunoassay (RIA). The method is routinely used in rodent endocrine and toxicity studies. The ELISA method is a commonly used as a human clinical test method. Analytical determination of iodothyronines (T3, T4, rT3, T2) and their conjugates, though methods employing HLPC, liquid chromatography, immuno luminescence, and mass spectrometry are less common, but are becoming increasingly available (Hornung et al., 2015; DeVito et al., 1999; Baret and Fert, 1989; Spencer, 2013).  It is important to note that thyroid hormones concentrations can be influenced by a number of intrinsic and extrinsic factors (e.g., circadian rhythms, stress, food intake, housing, noise) (see for example, Döhler et al., 1979).

Any of these measurements should be evaluated for the relationship to the actual endpoint of interest, repeatability, reproducibility, and lower limits of quantification using a fit-for-purpose approach (i.e., different regulatory needs will require different levels of confidence in the AOP). This is of particular significance when assessing the very low levels of TH present in fetal serum. Detection limits of the assay must be compatible with the levels in the biological sample.  All three of the methods summarized above would be fit-for-purpose, depending on the number of samples to be evaluated and the associated costs of each method. Both RIA and ELISA measure THs by an indirect methodology, whereas analytical determination is the most direct measurement available. All these methods, particularly RIA, are repeatable and reproducible.

 

References

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Baret A. and Fert V.  T4 and ultrasensitive TSH immunoassays using luminescent enhanced xanthine oxidase assay. J Biolumin Chemilumin. 1989. 4(1):149-153

Bartalena L, Robbins J. Thyroid hormone transport proteins. Clin Lab Med. 1993 Sep;13(3):583-98. Bassett JH, Harvey CB, Williams GR. (2003). Mechanisms of thyroid hormone receptor-specific nuclear and extra nuclear actions. Mol Cell Endocrinol. 213:1-11.

Capen CC. Mechanistic data and risk assessment of selected toxic end points of the thyroid gland. Toxicol Pathol. 1997 25(1):39-48.

Cope RB, Kacew S, Dourson M. A reproductive, developmental and neurobehavioral study following oral exposure of tetrabromobisphenol A on Sprague-Dawley rats. Toxicology. 2015 329:49-59.

Corcoran JM, Eastman CJ, Carter JN, Lazarus L. (1977). Circulating thyroid hormone levels in children. Arch Dis Child. 52: 716-720.

Crofton KM. Developmental disruption of thyroid hormone: correlations with hearing dysfunction in rats. Risk Anal. 2004 Dec;24(6):1665-71.

DeVito M, Biegel L, Brouwer A, Brown S, Brucker-Davis F, Cheek AO, Christensen R, Colborn T, Cooke P, Crissman J, Crofton K, Doerge D, Gray E, Hauser P, Hurley P, Kohn M, Lazar J, McMaster S, McClain M, McConnell E, Meier C, Miller R, Tietge J, Tyl R. (1999). Screening methods for thyroid hormone disruptors. Environ Health Perspect. 107:407-415.

Döhler KD, Wong CC, von zur Mühlen A (1979).   The rat as model for the study of drug effects on thyroid function: consideration of methodological problems.  Pharmacol Ther B. 5:305-18.

Eales JG. (1997). Iodine metabolism and thyroid related functions in organisms lacking thyroid follicles: Are thyroid hormones also vitaminsProc Soc Exp Biol Med. 214:302-317.

Furlow JD, Neff ES. (2006). A developmental switch induced by thyroid hormone: Xenopus laevis metamorphosis. Trends Endocrinol Metab. 17:40–47.

Goldey ES, Crofton KM. Thyroxine replacement attenuates hypothyroxinemia, hearing loss, and motor deficits following developmental exposure to Aroclor 1254 in rats. Toxicol Sci. 1998 45(1):94-10

Goldey ES, Kehn LS, Lau C, Rehnberg GL, Crofton KM.  Developmental exposure to polychlorinated biphenyls (Aroclor 1254) reduces circulating thyroid hormone concentrations and causes hearing deficits in rats. Tox Appl Pharmacol. 1995 135(1):77-88.

Harris AR, Fang SL, Prosky J, Braverman LE, Vagenakis AG.  Decreased outer ring monodeiodination of thyroxine and reverse triiodothyronine in the fetal and neonatal rat.  Endocrinology. 1978 Dec;103(6):2216-22

Heyland A, Hodin J. (2004). Heterochronic developmental shift caused by thyroid hormone in larval sand dollars and its implications for phenotypic plasticity and the evolution of non-feeding development. Evolution. 58: 524-538.

Heyland A, Moroz LL. (2005). Cross-kingdom hormonal signaling: an insight from thyroid hormone functions in marine larvae. J Exp Biol. 208:4355-4361.

 Hill RN, Crisp TM, Hurley PM, Rosenthal SL, Singh DV. Risk assessment of thyroid follicular cell tumors.  Environ Health Perspect. 1998 Aug;106(8):447-57.

Hornung MW, Kosian P, Haselman J, Korte J, Challis K, Macherla C, Nevalainen E, Degitz S (2015) In vitro, ex vivo and in vivo determination of thyroid hormone modulating activity of benzothiazoles. Toxicol Sci 146:254-264.

Hulbert AJ. Thyroid hormones and their effects: a new perspective. Biol Rev Camb Philos Soc. 2000 Nov;75(4):519-631. Review.

Kapelari K, Kirchlechner C, Högler W, Schweitzer K, Virgolini I, Moncayo R. 2008. Pediatric reference intervals for thyroid hormone levels from birth to adulthood: a retrospective study. BMC Endocr Disord. 8: 15.

Lau C, Thibodeaux JR, Hanson RG, Rogers JM, Grey BE, Stanton ME, Butenhoff JL, Stevenson LA.  Exposure to perfluorooctane sulfonate during pregnancy in rat and mouse. II: postnatal evaluation.  Toxicol Sci. 2003 Aug;74(2):382-92.

Leloup, J., and M. Buscaglia. La triiodothyronine: hormone de la métamorphose des amphibiens. CR Acad Sci 284 (1977): 2261-2263.

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McClain RM. Mechanistic considerations for the relevance of animal data on thyroid neoplasia to human risk assessment. Mutat Res. 1995 Dec;333(1-2):131-42

Miller MD, Crofton KM, Rice DC, Zoeller RT.  Thyroid-disrupting chemicals: interpreting upstream biomarkers of adverse outcomes. Environ Health Perspect. 2009 117(7):1033-41

Morse DC, Wehler EK, Wesseling W, Koeman JH, Brouwer A. Alterations in rat brain thyroid hormone status following pre- and postnatal exposure to polychlorinated biphenyls (Aroclor 1254). Toxicol Appl Pharmacol. 1996 Feb;136(2):269-79.

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Key Event Component

Key Event Component

Key Event Component

Domain of Applicability

Key Event Description

The hippocampus is a major brain region located in the medial temporal lobe in humans and other mammals (West, 1990). Developmentally it is derived from neuronal and glial cells in the neural tube and differentiates in the proencephalon and telencephalon.  The hippocampus is a cortical structure, but only contains 3-layers, distinct from the 6-layered neocortical structures. For this reason, it is known as archicortex or paleocortex meaning old cortex. Within humans, the structure is identified as early as fetal week 13 and matures rapidly until 2 to 3 years of age (Kier et al 1997), with continuing slow growth thereafter until adult ages (Utsunomiya et al., 1999).  In rodents, the hippocampus begins to form in midgestation, with the CA fields forming in advance of the dentate gyrus. Dentate gyrus forms in late gestation with most of its development occurring in the first 2-3 postnatal weeks (Altman and Bayer, 1990a; 1990b).

The structure of the hippocampus has been divided into regions that include CA1 through CA4 and the dentate gyrus. The principal cell bodies of the CA field are pyramidal neurons, those of the dentate gyrus are granule cells. The dentate gyrus forms later in development than the CA fields of the hippocampus. These regions are generally found in all mammalian hippocampi.

The major input pathway to the hippocampus is from the layer 2 neurons of the entorhinal cortex to the dentate gyrus via the perforant path forming the first connection of the trisynaptic loop of the hippocampal circuit. Direct afferents from the dentate gyrus (mossy fibers) then synapse on CA3 pyramidal cells which in turn send their axons (Schaeffer Collaterals) to CA1 neurons to complete the trisynaptic circuit (Figure 1). From the CA fields information then passes through the subiculum entering the fiber pathways of the alveus, fimbria, and fornix and it routed to other areas of the brain (Amaral and Lavenex, 2006). Through the interconnectivity within the hippocampus and its connections to amygdala, septum and cortex, the hippocampus plays a pivotal role in several learning and memory processes, including spatial behaviors. The primary input pathway to the CA regions of the hippocampus is from the septum by way of the fornix and direct input from the amygdala. Reciprocal outputs from the hippocampus back to these regions and beyond also exist.

 

Trisynaptic Hippocampal Circuitry

How it is Measured or Detected

Data in support of this key event have been collected using a wide variety of standard biochemical, histological and anatomical methods (e.g., morphometrics, immunohistochemical staining, in situ hybridization and imaging procedures). Many of methods applied to reveal anatomical abnormalities are routine neurohistopathology procedures similar to those recommended in EPA and OECD developmental neurotoxicity guidelines (US EPA, 1998; OCED, 2007). Subtle cytoarchitectural features depend on more specialized birth dating procedures and staining techniques. It is essential to consider the timing of events during development for detection to occur, as well as the timing for detection (Hevner, 2007; Garman et al., 2001; Zgraggen et al., 2012). Similar techniques used in rodent stydies have been applied to postmortem tissue in humans. 

In humans, structural neuroimaging techniques are used to assess hippocampal volume with an analysis technique known as voxel-based morphometry (VBM). Volume of brain regions is measured by drawing regions of interest (ROIs) on images from brain scans obtained from magnetic resonance imaging (MRI) or positron emission tomography (PET) scans and calculating the volume enclosed. (Mechelli et al., 2005). Similar imaging techniques can be applied in rodent models (Powell et al., 2009; Hasegawa et al., 2010; Pirko et al., 2005; Pirko and Johnson, 2008).

References

Altman J, Bayer SA. Migration and distribution of two populations of hippocampal granule cell precursors during the perinatal and postnatal periods. J Comp Neurol. 1990a Nov 15;301(3):365-81.

Altman J, Bayer SA. Prolonged sojourn of developing pyramidal cells in the intermediate zone of the hippocampus and their settling in the stratum pyramidale. J Comp Neurol. 1990b Nov 15;301(3):343-64.

Amaral D, Lavenex P (2006). "Ch 3. Hippocampal Neuroanatomy". In Andersen P, Morris R, Amaral D, Bliss T, O'Keefe J. The Hippocampus Book. Oxford University Press. ISBN 978-0-19-510027-3.

Garman RH, Fix AS, Jortner BS, Jensen KF, Hardisty JF, Claudio L, Ferenc S. Methods to identify and characterize developmental neurotoxicity for human health risk assessment. II: neuropathology. Environ Health Perspect. 2001 Mar;109 Suppl 1:93-100.

Hasegawa M, Kida I, Wada H.  A volumetric analysis of the brain and hippocampus of rats rendered perinatal hypothyroid. Neurosci Lett. 2010 Aug 2;479(3):240-4.

Hevner RF. Layer-specific markers as probes for neuron type identity in human neocortex and malformations of cortical development. J Neuropathol Exp Neurol. 2007 66(2):101-9.

Kier, EL, Kim, JH, Fulbright, K, Bronen, RA. Embryology of the human fetal hippocampus: MR imaging, anatomy, and histology. AJNR Am J Neuroradiol: 1997, 18(3);525-32.

Mechelli A, Price C, Friston K, Ashburner J (2005) Voxel-Based Morphometry of the Human Brain: Methods and Applications. Curr Med Imaging Rev 1:105-113.

OECD. 2007. OECD guidelines for the testing of chemicals/ section 4: Health effects. Test no. 426: Developmental neurotoxicity study. http://www.oecd.org/dataoecd/20/52/37622194.

Pirko I, Fricke ST, Johnson AJ, Rodriguez M, Macura SI. Magnetic resonance imaging, microscopy, and spectroscopy of the central nervous system in experimental animals. NeuroRx. 2005 Apr;2(2):250-64.

Pirko I, Johnson AJ. Neuroimaging of demyelination and remyelination models. Curr Top Microbiol Immunol. 2008; 318:241-66.

Powell MH, Nguyen HV, Gilbert M, Parekh M, Colon-Perez LM, Mareci TH, Montie E. Magnetic resonance imaging and volumetric analysis: novel tools to study the effects of thyroid hormone disruption on white matter development. Neurotoxicology. 2012 Oct;33(5):1322-9.

U.S.EPA. 1998. Health effects guidelines OPPTS 870.6300 developmental neurotoxicity study. EPA Document 712-C-98-239.Office of Prevention Pesticides and Toxic Substances.

Utsunomiya, H., K Takano, M Okazaki, A Mitsudome Development of the temporal lobe in infants and children: analysis by MR-based volumetry. AJNR Am J Neuroradiol: 1999, 20(4);717-23.

West MJ (1990). "Stereological studies of the hippocampus: a comparison of the hippocampal subdivisions of diverse species including hedgehogs, laboratory rodents, wild mice and men". Progress in Brain Research. Progress in Brain Research 83: 13–36.

Zgraggen E, Boitard M, Roman I, Kanemitsu M, Potter G, Salmon P, Vutskits L, Dayer AG, Kiss JZ. Early postnatal migration and development of layer II pyramidal neurons in the rodent cingulate/retrosplenial cortex. Cereb Cortex. 2012 Jan;22(1):144-57. 

 

Key Event Component

Domain of Applicability

Key Event Description

The hippocampus functions as a highly integrated and organized communication and information processing network with millions of interconnections among its constitutive neurons. Neurons in the hippocampus and throughout the brain transmit and receive information largely through chemical transmission across the synaptic cleft, the space where the specialized ending of the presynaptic axon terminus of the transmitting neuron meets the specialized postsynaptic region of the neuron that is receiving that information (Kandell et al., 2012).

During development (see KE: Hippocampal anatomy, Altered), as neurons reach their final destination and extend axonal processes, early patterns of electrical synaptic activity emerge in the hippocampus. These are large fields of axonal innervation of broad synaptic target sites that are replaced by more elaborate but highly targeted and refined axonal projections brought about by activity-dependent synaptic pruning and synapse elimination.  This is a classic case of the interaction between physiological and anatomical development, where anatomy develops first, and can be ‘reshaped’ by physiological function (Kutsarova et al., 2017).

In the rat, excitatory processes are fully mature in area CA1 of hippocampus within 2 weeks of birth with inhibitory processes lagging begin by several weeks (Muller et al., 1989; Michelson and Lothman, 1988; Harris and Teyler, 1984). In hippocampal slices, inhibitory function in areaCA1s is first seen on postnatal day 5 and increases in strength at postnatal day 12 through 15.  In vivo studies fail to detect inhibition until postnatal day 18 with steady increase thereafter to adult levels by postnatal day 28. Synaptic plasticity in the form of long-term potentiation (LTP) is absent in the very young animal, only emerging about postnatal day 14, appearing to require the stability of both excitatory and inhibitory function to be established (Muller et al., 1989; Bekenstein and Lothman, 1991). These features of the maturation of hippocampal physiology are paralleled in dentate gyrus, but as with anatomical indices in the rat, the development of these physiological parameters lag behind the CA1 by about 1 week. 

How it is Measured or Detected

In animals, synaptic function in the hippocampus has been examined with imaging techniques, but more routinely, electrical field potentials recorded in two subregions of the hippocampus, area CA1 and dentate gyrus, have been assessed in vivo or in vitro from slices taken from naive or exposed animals. Field potentials reflect the summed synaptic response of a population of neurons following direct stimulation of input pathways across a monosynaptic connection. Changes in response amplitude due to chemical perturbations and other stressors (e.g., iodine deficiency, thyroidectomy, gene knockouts) is evidence of altered synaptic function. This can be measured in vitro, in vivo, or in hippocampal slices taken from treated animals (Gilbert and Burdette, 1995). The most common physiological measurements used to assess function of the hippocampus are excitatory synaptic transmission, inhibitory synaptic transmission, and synaptic plasticity in the form of long-term potentiation (LTP).

Excitatory Synaptic Transmission: Two measures, the excitatory postsynaptic potential (EPSP) and the population spike are derived from the compound field potential at increasing stimulus strengths. The function described by the relationship of current strength (input, I) and evoked response (output, O), the I-O curve is the measure of excitatory synaptic transmission (Gilbert and Burdette, 1995).

Inhibitory Synaptic Transmission: Pairs of stimulus pulses delivered in close temporal proximity is used to probe the integrity of inhibitory synaptic transmission. The response evoked by the second pulse of the pair at brief intervals (<30 msec) arrives during the activation of feedback inhibitory loops in the hippocampus. An alteration in the degree of suppression to the 2nd pulse of the pair reflects altered inhibitory synaptic function (Gilbert and Burdette, 1995).

Long Term Potentiation (LTP): LTP is widely accepted to be a major component of the cellular processes that underlie learning and memory (Malenka and Bear, 2004; Bramham and Messaoudi, 2005). LTP represents, at the synapse and molecular level, the coincident firing of large numbers of neurons that are engaged during a learning event. The persistence of LTP emulates the duration of the memory. Synaptic plasticity in the form of LTP is assessed by delivering trains of high frequency stimulation to induce a prolonged augmentation of synaptic response. Probe stimuli at midrange stimulus strengths are delivered before and after application of LTP-inducing trains. The degree of increase in EPSP and PS amplitude to the probe stimulus after train application, and the duration of the induced synaptic enhancement are metrics of LTP. Additionally, contrasting I-O functions of excitatory synaptic transmission before and after (hours to days) LTP is induced is also a common measure of LTP maintanence (Bramham and Messaoudi, 2005; Kandell et al., 2012; Malenka and Bear, 2004).

Synaptic function in the human hippocampus has been assessed using electroencephalography (EEG) and functional neuroimaging techniques (Clapp et al., 2012). EEG is a measure of electrical activity over many brain regions but primarily from the cortex using small flat metal discs (electrodes) placed over the surface of the skull. It is a readily available test that provides evidence of how the brain functions over time. Functional magnetic resonance imaging or functional MRI (fMRI) uses MRI technology to measure brain activity by detecting associated changes in blood flow. This technique relies on the fact that cerebral blood flow and neuronal activation are coupled. Positron emission tomography (PET) is a functional imaging technique that detects pairs of gamma rays emitted indirectly by a radionuclide (tracer) injected into the body (Tietze, 2012; McCarthy, 1995). Like fMRI, PET scans indirectly measure blood flow to different parts of the brain – the higher the blood flow, the greater the activation (McCarthy, 1995). These techniques have been widely applied in clinical and research settings to assess learning and memory in humans and can provide information targeted to hippocampal functionality (McCarthy, 1995; Smith and Jonides, 1997; Willoughby et al., 2014; Wheeler et al., 2015; Gilbert et al., 1998).

Assays of this type are fit for purpose, have been well accepted in the literature, and are reproducible across laboratories. The assay directly measures the key event of altered neurophysiological function.

References

Bekenstein JW, Lothman EW. An in vivo study of the ontogeny of long-term potentiation (LTP) in the CA1 region and in the dentate gyrus of the rat hippocampal formation. Brain Res Dev Brain Res. 1991 Nov 19;63(1-2):245-

Bramham CR, Messaoudi E (2005) BDNF function in adult synaptic plasticity: the synaptic consolidation hypothesis. Prog Neurobiol 76:99-125.

Clapp WC, Hamm JP, Kirk IJ, Teyler TJ. Translating long-term potentiation from animals to humans: a novel method for noninvasive assessment of cortical plasticity. Biol Psychiatry. 2012 Mar 15;71(6):496-502.

Gilbert, M.E. and Burdette, L.J. (1995). Hippocampal Field Potentials: A Model System to Characterize Neurotoxicity. In Neurotoxicology: Approaches and Methods. L.W Chang and W. Slikker (Eds). Academic Press:New York, 183-204.

Gilbert ME, Mack CM. Chronic lead exposure accelerates decay of long-term potentiation in rat dentate gyrus in vivo. Brain Res. 1998 Apr 6;789(1):139-49.

Harris KM, Teyler TJ. Developmental onset of long-term potentiation in area CA1 of the rat hippocampus. J Physiol. 1984. 346:27-48.

Kandell, E., Schwartz, J., Siegelbaum, A. and Hudspeth, A.J.  (2012) Principles of Neural Science, 5^th Edition.  Elsevier, North Holland.

Kutsarova E, Munz M, Ruthazer ES.  Rules for Shaping Neural Connections in the Developing Brain.  Front Neural Circuits. 2017 Jan 10;10:111. doi: 10.3389/fncir.2016.00111.

Malenka RC, Bear MF (2004) LTP and LTD: an embarrassment of riches. Neuron 44:5-21.

McCarthy, G. (1995) Review: Functional Neuroimaging and Memory. The Neuroscientist, 1:155-163.

Michelson HB, Lothman EW. An in vivo electrophysiological study of the ontogeny of excitatory and inhibitory processes in the rat hippocampus. Brain Res Dev Brain Res. 1989 May 1;47(1):113-22.

Muller D, Oliver M, Lynch G. Developmental changes in synaptic properties in hippocampus of neonatal rats. Brain Res Dev Brain Res. 1989 Sep 1;49(1):105-14.

Smith, E and Jonides, J. (1997). Working Memory: A View from Neuroimaging. Cognitive Psychology, 33:5-42.

Tietze, KJ. (2012). Review of Laboratory and Diagnostic Tests- Positron Emission Tomography. In Clinical Sills for Pharmacists, 3rd Edition, pp 86-122. 

Wheeler SM, McLelland VC, Sheard E, McAndrews MP, Rovet JF (2015) Hippocampal Functioning and Verbal Associative Memory in Adolescents with Congenital Hypothyroidism. Front Endocrinol (Lausanne) 6:163.

Willoughby KA, McAndrews MP, Rovet JF (2014) Effects of maternal hypothyroidism on offspring hippocampus and memory. Thyroid 24:576-584.

Key Event Component

Domain of Applicability

Key Event Description

Thyroid hormones control genes in the developing brain by classical ligand (T3) activation of thyroid receptors which leads to DNA binding and subsequent transcription and translation (for a review of TH rols in brain development see, Bernal 2015). Gene expression profiles have been published for the developing human and rodent hippocampus (Zhang et al., 2002; Mody et al., 2001). In both humans and rodents, the hippocampus undergoes typical stages of neurodevelopment found in most brain regions, including: cell proliferation, migration, differentiation, synapse formation, and the maturation of synaptic function. In the rodent, peak windows during pre- and post-natal periods have been identified during which major cellular and physiological events occur (see Figure 1). Each window expresses distinct patterns of gene transcription and clusters of genes increase their expression corresponding to the progression of events of hippocampal ontogeny (see Mody et al., 2001).  Tables of gene clusters associated with these phases can be found in Supplementary Tables of Mody et al. (2001).

 

During the very early prenatal period, genes corresponding to general cellular function are prominent (Mody et al., 2001).  These are followed in time by genes regulating neuronal differentiation and migration in the mid to late gestational period. From late gestation (gestational day 15) until birth almost all the cells in the CA fields switch from a highly active proliferation state to a postmitotic state, and then undergo differentiation and migration. Expression of proliferative genes involved in cell cycle progression are highly expressed at gestational day 16, then subsequently are silent immediately after birth when genes directing neuronal growth switch on. The pyramidal neurons of the CA fields in the hippocampus proper develop in advance of the granule cells that comprise the principal cells of the dentate gyrus. As such, the genes controlling the distinct phases of neurodevelopment are expressed at different times in these two hippocampal subregions (Altman and Bayer, 1990a; b). In both subregions, however, many phenotypic changes within the hippocampal neuron occur in the period immediately after birth (postnatal day 1 to 7). Almost all neurons show extensive growth and differentiation during the first postnatal week. These cellular changes are marked by rapid cytoskeletal changes, production of cell adhesion molecules, and extracellular matrix formation. The gene families involved in these processes include actins, tubulins, and chaperonin proteins essential for promoting correct protein folding of cytoskeletal components. Cell adhesion and extracellular matrix proteins are also upregulated during this period as these genes are critical for differentiation and synaptogenesis.

During late postnatal hippocampal development (postnatal day 16-30), hippocampal circuits become more active and exhibit increased synaptic plasticity. Many genes upregulated during this phase of development are involved in synaptic function and include genes regulating vesicle associated proteins and calcium-mediated transmitter release, neurotrophins, and neurotransmitter receptors. Efficient energy utilization is essential during this period of increased synaptic activity, events mirrored by an upregulation of enzymes involved in glucose and oxidative metabolism.

How it is Measured or Detected

Measurement of genomic profiles in developing brain use methods that are well established and accepted in the published literature.  Microarray studies with expression profile analyses have been conducted in cortex and hippocampus of humans (Zhang et al., 2002), non-human primates, and rodent brains of various ages (Mody et al., 2001; Royland et al., 2008; Dong et al., 2015). More commonly, quantitative rtPCR or in situ hybridization have been used to probe individual gene transcripts (Dowling et al., 2000, Morte et al., 2010) or their protein products (Alvarez-Dolado et al., 1994; Gilbert et al., 2007). Recently RNA-Seq technology was applied to T3-treated primary mouse cortical cells and gene targets enriched in astrocytes and neurons to identify TH-responsive genes (Gil-Ibanez et al, 2015).

References

Altman J, Bayer SA. Migration and distribution of two populations of hippocampal granule cell precursors during the perinatal and postnatal periods. J Comp Neurol. 1990a Nov 15;301(3):365-81.

Altman J, Bayer SA. Prolonged sojourn of developing pyramidal cells in the intermediate zone of the hippocampus and their settling in the stratum pyramidale. J Comp Neurol. 1990b Nov 15;301(3):343-64.

Alvarez-Dolado M, Ruiz M, Del Rio JA, Alcantara S, Burgaya F, Sheldon M, Nakajima K, Bernal J, Howell BW, Curran T, Soriano E, Munoz A (1999) Thyroid hormone regulates reelin and dab1 expression during brain development. J Neurosci 19:6979-6993.

Bernal J. (2105)  Thyroid Hormones in Brain Development and Function. In: De Groot LJ, Chrousos G, Dungan K, Feingold KR, Grossman A, Hershman JM, Koch C, Korbonits M, McLachlan R, New M, Purnell J, Rebar R, Singer F, Vinik A, editors. Endotext [Internet]. South Dartmouth (MA): MDText.com, Inc..

Dong H, You SH, Williams A, Wade MG, Yauk CL, Thomas Zoeller R (2015) Transient Maternal Hypothyroxinemia Potentiates the Transcriptional Response to Exogenous Thyroid Hormone in the Fetal Cerebral Cortex Before the Onset of Fetal Thyroid Function: A Messenger and MicroRNA Profiling Study. Cereb Cortex 25:1735-1745.

Dowling AL, Zoeller RT. 2000. Thyroid hormone of maternal origin regulates the expression of RC3/neurogranin mRNA in the fetal rat brain. Brain Res: Molec Brain Res.  82:126-132.

Gilbert ME, Sui L, Walker MJ, Anderson W, Thomas S, Smoller SN, Schon JP, Phani S, Goodman JH (2007) Thyroid hormone insufficiency during brain development reduces parvalbumin immunoreactivity and inhibitory function in the hippocampus. Endocrinology 148:92-102.

Gil-Ibanez P, Garcia-Garcia F, Dopazo J, Bernal J, Morte B. 2015. Global Transcriptome Analysis of Primary Cerebrocortical Cells: Identification of Genes Regulated by Triiodothyronine in Specific Cell Types. Cerebral cortex. Nov 2.

Kempermann G.  New neurons for 'survival of the fittest'.  Nat Rev Neurosci. 2012 Oct;13(10):727-36.

Mody M, Cao Y, Cui Z, Tay KY, Shyong A, Shimizu E, Pham K, Schultz P, Welsh D, Tsien JZ. Genome-wide gene expression profiles of the developing mouse hippocampus. Proc Natl Acad Sci U S A. 2001 Jul 17;98(15):8862-7.

Morte B, Ceballos A, Diez D, Grijota-Martinez C, Dumitrescu AM, Di Cosmo C, Galton VA, Refetoff S, Bernal J.  Thyroid hormone-regulated mouse cerebral cortex genes are differentially dependent on the source of the hormone: a study in monocarboxylate transporter-8- and deiodinase-2-deficient mice. Endocrinology. 2010. 151:2381-2387.

Royland JE, Parker JS, Gilbert ME. A genomic analysis of subclinical hypothyroidism in hippocampus and neocortex of the developing rat brain. J Neuroendocrinol. 2008 Dec;20(12):1319-38.

Zhang Y, Mei P, Lou R, Zhang MQ, Wu G, Qiang B, Zhang Z, Shen Y. Gene expression profiling in developing human hippocampus. J Neurosci Res. 2002 Oct 15;70(2):200-8.

Key Event Component