page 1 of 48 diabetes...1 thyroid hormone coordinates pancreatic islet maturation during the...

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Thyroid hormone coordinates pancreatic islet maturation during the zebrafish larval to

juvenile transition to maintain glucose homeostasis

Hiroki Matsuda1*

, Sri Teja Mullapudi1, Yuxi Zhang

2, Daniel Hesselson

2,3, and Didier Y. R.

Stainier1*

Author affiliation:

1. Max Planck Institute for Heart and Lung Research, Department of Developmental

Genetics, Bad Nauheim, Germany

2. Garvan Institute of Medical Research, Diabetes and Metabolism Division, Sydney,

Australia

3. St. Vincent’s Clinical School, UNSW Australia, Sydney, Australia

*Corresponding authors

Hiroki Matsuda ([email protected])

Didier Stainier ([email protected])

Department of Developmental Genetics

Max Planck Institute for Heart and Lung Research.

Ludwigstrasse 43, 61231 Bad Nauheim, Germany

Phone: +49 (0) 6032 705-1333

Fax:+49 (0) 6032 705-1304

Running title: TH regulates pancreatic islet maturation

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Diabetes Publish Ahead of Print, published online July 11, 2017

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Abstract

Thyroid hormone (TH) signaling promotes tissue maturation and adult organ

formation. Developmental transitions alter an organism's metabolic requirements, and it

remains unclear how development and metabolic demands are coordinated. We used the

zebrafish as a model to test whether and how TH signaling affects pancreatic islet maturation,

and consequently glucose homeostasis, during the larval to juvenile transition. We found that

exogenous TH precociously activates the β cell differentiation genes pax6b and mnx1, while

downregulating arxa, a master regulator of α cell development and function. Together these

effects induced hypoglycemia, at least in part by increasing insulin and decreasing glucagon

expression. We visualized TH target tissues using a novel TH responsive reporter line and

found that both α and β cells become targets of endogenous TH signaling during the larval to

juvenile transition. Importantly, endogenous TH is required during this transition for the

functional maturation of α and β cells in order to maintain glucose homeostasis. Thus, our

study sheds new light on the regulation of glucose metabolism during major developmental

transitions.

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Introduction

Organ development often involves a two-step process: the formation of a functionally

immature organ during embryogenesis followed by maturation into the adult form. This

second step most often takes place during the postembryonic/postnatal developmental period,

when plasma thyroid hormone (TH) concentrations are high. In rodents, these major

developmental changes occur during the suckling to weaning transition, and their digestive

organs undergo functional and morphological changes during this period (1), correlating with

the dietary switch from mother’s milk to solid foods. It has been proposed that Xenopus

metamorphosis and the zebrafish larval-juvenile transition are functionally equivalent to

mammalian weaning, based on cellular and molecular events in their digestive organs during

these periods (2-7).

The pancreas is an organ in the digestive system that undergoes morphological and

functional changes during the suckling to weaning transition. For example, the weight of the

rat pancreas increases 5-6 fold during this period (8). In addition, pancreatic acinar cells

become functionally mature and start to secrete the full suite of digestive enzymes, including

trypsin and amylase, into the duodenum during the weaning period (1). In the endocrine

pancreas, β cell mass increases (8), and more organized, functional islets appear during this

period (8-13) as β cells acquire the ability to secrete insulin in response to glucose around the

second week after birth (9). β cell secretory functions continue to improve well beyond

weaning periods (9), although the underlying mechanisms are not well understood.

The thyroid hormone T3 (triiodothyronine) signals through the nuclear Thyroid

Hormone Receptor (THR). THR functions as a transcriptional repressor in the absence of TH

and a transcriptional activator in the presence of TH. It has been reported that ligand bound

THR can directly activate Mafa expression and that exogenous TH can induce MAFA

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dependent glucose-responsive insulin secretion in the rat pancreas (14). Furthermore, TH is

used in the most effective protocols developed to date to generate glucose responsive insulin

secreting cells from pluripotent cells (15, 16). These data implicate TH signaling in β cell

maturation. Furthermore, TH can also induce endocrine cell differentiation from ductal cells

in E12.5 mouse pancreatic explants (17) and stimulate acinar cell proliferation in the adult

mouse (18). It has therefore been hypothesized that TH regulates multiple aspects of

pancreas development, including endocrine cell differentiation, β cell maturation, and the size

of the exocrine compartment. However, it remains unclear exactly when and where TH

signaling is activated during pancreas development.

The zebrafish (Danio rerio) has emerged as a powerful model to study mechanisms of

pancreas development and β cell differentiation. Zebrafish pancreatic development is similar

to that in mammals in terms of cellular events and molecular pathways (19-21). For example,

in zebrafish, bilateral pdx1 positive pancreatic primordia appear by 14 hours post fertilization

(hpf), and the dorsal bud emerges from these primordia by 22 hpf. At 40 hpf, the ventral bud

is visible, and by 52 hpf, it has fused with the dorsal bud; the principal islet, originally

derived from the dorsal bud, is organized like a mammalian islet, with α and δ cells

surrounding a β cell core. Here we investigated the spatiotemporal regulation of TH

signaling during zebrafish pancreatic development, focusing on the endocrine lineages. We

show that TH stimulates insulin expression, through upregulation of pax6b and mnx1. In

addition, we found that TH inhibits the expression of both glucagon paralogs by upregulating

pax4 expression and downregulating arxa expression. Together these effects, at least in part,

decrease glucose levels in zebrafish larvae. To visualize TH target tissues, we generated a

TH reporter line and found that both α and β cells are direct targets of TH signaling during

and after the larval-juvenile transition. Additionally, using a hypothyroid zebrafish model,

we found that TH can regulate glucose homeostasis by modulating the expression of

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endocrine genes during the larval-juvenile transition. In adults, TH continues to regulate α

and β cell function as well as glucose homeostasis. These results indicate that TH signaling

is involved in the development and maturation of α and β cells during postembryonic

development, and that it maintains glucose homeostasis, at least in part, by modulating the

insulin to glucagon ratio throughout the vertebrate lifecycle.

Research Design and Methods

Zebrafish lines

All zebrafish husbandry was performed under standard conditions in accordance with

institutional and national ethical and animal welfare guidelines. A 1000x stock solution of 10

µM triiodothyronine (T3) in 5 mM NaOH was made and stored at – 20 ºC until use. Larvae

were treated with the indicated concentration of T3 in egg water for the indicated number of

days at 28°C. Larvae incubated in 5 µM NaOH egg water were used as controls. For adult

zebrafish, 10 µl of 10 nM T3 was injected intraperitoneally once a day for three consecutive

days. 10 µl of 7 µM NaOH in egg water were used for control injections.

We used the following transgenic lines: Tg(ins:Luc2;cryaa:mCherry)gi3

, abbreviated

ins:Luc2; Tg(ins:H2BGFP;ins:dsRED)s960

(22), abbreviated ins:H2BGFP; Tg(-

4.0ins:eGFP)Zf5

(23), abbreviated ins:eGFP; Tg(gcga:eGFP)ia1

(24), abbreviated gcga:eGFP;

Tg(P0-pax6b:eGFP)ulg515

(25), abbreviated pax6b:eGFP; TgBAC(neurod1:eGFP)nl1

(26),

abbreviated neurod1:eGFP; Tg(-2.6mnx1:GFP)ml59

(27), abbreviated mnx1:eGFP;

Tg(tg:nVenus-2a-nfsB)wp.rt8

(28), abbreviated tg:nVenus-2a-nfsB.

Nitroreductase-mediated cell ablation

To ablate the thyroid follicles of tg:nVenus-2a-nfsB fish, we incubated 20 days post

fertilization (dpf) larvae for 48 h in 10 mM Metronidazole (Mtz) with 1% DMSO, or 1%

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DMSO alone as a control. Thyroid ablations of adult fish were performed on transgenic fish

injected intraperitoneally with 15 µl of 30 mM Mtz or DMSO as a control.

Results

T3 stimulates insulin promoter activity, reduces glucose levels and improves glucose

tolerance

To determine the effect of thyroid hormone on pancreatic preproinsulin (insulin)

expression, transgenic insulin:luciferase2 (Tg(ins:Lyc2;cryaa:mCherry)gi3

) larvae were

treated with T3 from 4 to 7 dpf (Fig. 1A and B). T3 significantly increased insulin promoter

activity at 10 nM, but showed no significant effects at 0.1, 1 or 100 nM (Fig. 1B). Glucose

levels were reciprocally reduced in a T3 dose-dependent manner (Fig. 1C). During the 4 to 7

day developmental window, 10 nM T3 was the optimal concentration for activation of insulin

expression and glucose reduction. To further define when T3 acts, larvae were treated with

10 nM T3 starting at 4 dpf and insulin promoter activity and glucose levels were analyzed at

5, 6, 7 and 8 dpf (Fig. 1D). Insulin promoter activity was enhanced at 7 and 8 dpf (Fig. 1E),

while glucose levels were reduced at 6,7 and 8 dpf (Fig. 1F). These results indicate that

insulin expression is T3 responsive by 7 dpf and that our 4 to 7 dpf treatment conditions

induce a hypoglycemic phenotype.

Given the correlation between insulin upregulation and glucose reduction in these

experiments, we next tested glucose tolerance in T3 treated larvae. At 7 dpf, after 3 days of

T3 treatment, larvae were exposed to glucose for 60 mins and glucose levels were analyzed at

0, 30, 60 and 90 mins after glucose treatment (Fig. 1G). Glucose levels were unaltered in T3

treated larvae immediately after the glucose challenge (0 min). Strikingly, at 30, 60 and 90

mins after the challenge, glucose levels were lower in T3-treated larvae compared to non-

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treated controls (Fig. 1H). These data indicate that T3 increases the glucose disposal capacity

of zebrafish larvae.

T3 stimulates insulin expression but does not alter β cell number

Given that TH stimulated insulin promoter activity in ins:luc2 lines, we next tested

whether TH stimulates endogenous insulin expression using qPCR. Zebrafish has two insulin

genes, preproinsulin (insulin) and preproinsulin b (insulin b). We focused our analysis on

insulin, because insulin b expression is minimal after 2 dpf. Starting T3 treatment at 4 dpf,

insulin expression was analyzed at 5, 6 and 7 dpf. Although T3 did not significantly affect

endogenous insulin expression at 5 and 6 dpf, enhanced insulin expression was observed at 7

dpf (Fig. 2A). Next, we analyzed whether TH alters the endogenous insulin expression

pattern using in situ hybridization. Zebrafish larvae were treated with T3 from 4 dpf and

analyzed at 7 dpf. T3 did not induce ectopic insulin expression (Fig. 2B and C). Furthermore,

using a similar treatment strategy with ins:eGFP reporter fish, we did not observe eGFP

expression outside the pancreas (Fig. 2D and E). Notably, T3 increased ins:eGFP expression

intensity in the principal islet compared to controls (Fig. 2F-H). To determine whether an

expanded β cell mass contributed to the increased insulin expression in T3-treated larvae, we

used a nuclear localized ins:H2BGFP reporter to quantify β cell number. Interestingly, T3

did not alter the number of β cells (Fig. 2I-K). Taken together, these data suggest that T3

treatments of zebrafish larvae enhance insulin expression without increasing β cell number in

zebrafish.

T3 represses glucagon expression and reduces α cell number

As glucose homeostasis is also under the control of Glucagon secreted from

pancreatic α cells, we investigated whether TH signaling regulates α cell function. We

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treated wild-type larvae with T3 starting at 4 dpf and analyzed the expression levels of

glucagon a (gcga) and glucagon b (gcgb) at 5, 6 and 7 dpf by qPCR. T3 significantly

repressed the expression of both gcga and gcgb at 5, 6 and 7 dpf (Fig. 3A). We next treated

larvae with T3 starting at 4 dpf and investigated its effect on the expression pattern of gcga

and gcgb at 7 dpf by in situ hybridization. T3 treatment reduced gcga and gcgb expression in

pancreatic islets (Fig. 3B) and no ectopic expression was observed (Fig. 3B and data not

shown). Consistent with the effect on endogenous gcga expression, T3 suppressed the

expression of the gcga:eGFP transcriptional reporter (Fig. 3C). Interestingly, the number of

gcga:eGFP positive α cells was modestly reduced in T3 treated larvae (26 +/- 2 in controls;

21+/- 2 in T3 treated, Fig. 3E). Thus, TH impairs α cell differentiation and/or proliferation in

addition to negatively regulating glucagon expression.

To determine whether the reduction in glucagon expression was caused by negative

feedback from elevated paracrine Insulin signaling upon T3 treatment, we generated insulin

(ins, previously known as insa) mutants using CRISPR/Cas9 mediated genome editing. ins-/-

larvae exhibit a complete absence of detectable Insulin protein in their pancreatic islet (data

not shown), and although they appear to develop apparently normally during early stages,

mutant animals die by around 10-11 dpf due to metabolic defects (data not shown). We

treated ins -/- larvae with T3 at 4 dpf and analyzed gcga and gcgb expression levels by qPCR

at 6 dpf. Both gcga and gcgb expression were reduced after T3 treatment (Supplemental

Figure 1). These results indicate that T3 represses glucagon expression through an Insulin-

independent pathway.

T3 stimulates pax6b, mnx1 and pax4 expression and represses arxa expression.

To understand how T3 regulates pancreatic islet expression of insulin, gcga and gcgb,

we investigated the effects of T3 treatment on upstream endocrine transcription factor genes.

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We used transcriptional eGFP reporters for pax6b, neurod1 and mnx1, which encode known

regulators of endocrine differentiation and/or maturation (Fig. 4A-F). T3 treatment increased

eGFP intensity of the pax6b and mnx1, but not of the neurod1, reporter transgenes within the

principal islet (Fig. 4A-F). Furthermore qPCR analysis revealed that T3 treatment

upregulated endogenous expression of pax6b, mnx1 and pax4, and downregulated expression

of arxa (Fig. 4G).

Pancreatic α and β cells are targets of thyroid hormone during and after larval

development.

We have shown that supplemental T3 stimulated insulin expression and repressed

glucagon expression leading to reduced glucose levels. We next asked whether and when

endogenous T3 signaling was activated in pancreatic α and β cells. To address this question,

we generated a TH responsive zebrafish line to visualize T3 target tissues in vivo. The

biological effects of T3 are mediated by signaling through THRs, which bind to TH response

elements (TRE) to regulate downstream target genes. Three copies of Xenopus laevis

THbzipTREs (6xTRE) were inserted upstream of candidate minimal promoters driving eGFP

to visualize TH signaling (Supplemental Figure 2A). To identify the optimal configuration,

four different minimal promoters (cfos, bglob1, tk and Pmini) were tested by transient

injection assay for their response to T3 treatment (Supplemental Figure 2B). The construct

incorporating the bglob1 minimal promoter showed the highest proportion of eGFP+

embryos after T3 treatment (Supplemental Figures 2C and D). Next, we confirmed that T3

responsiveness was mediated by the TREs. We compared 6xTRE to bglob1:eGFP plasmids

lacking TREs, containing 2xTREs, or containing mutant TREs that cannot bind THR

(6xTREs(mut)) (Supplemental Figure 2E), and found that intact TREs were essential for the

T3-dependent stimulation of 6xTRE-bglobe:eGFP expression (Supplemental Figure 2F).

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A stable transgenic 6xTRE-bglob1:eGFP reporter was used to identify cells

responsive to endogenous TH in the pancreas (Fig. 5A). First, to validate the reporter, we

confirmed that exogenous T3 induced ubiquitous eGFP expression during larval stages

(Supplemental Figures 3A, D and E). Before 20 dpf (Fig. 5B and G), including early larval

stages (Supplemental Figures 3B and C), eGFP positive cells were not detected in the

pancreas (Supplemental Figure 3C). By 25 dpf, eGFP expression was detected in 71 % of

Insulin positive cells (Fig. 5C and G, white arrows), but only 0.7 % of Glucagon positive

cells (red arrows). By 30 dpf, eGFP expression was also observed in 86 % of Glucagon

positive cells (Fig. 5D and G). In the adult pancreas, TH reporter expression was observed in

89 % of Insulin and 95 % of Glucagon positive cells (Fig. 5E and G). These results indicate

that endogenous TH signaling is active during and after the larval-juvenile transition in

differentiated α and β cells.

Thyroid hormone regulates glucose metabolism by stimulating insulin expression and

suppressing glucagon expression during the larval-juvenile transition.

Analysis of the TH reporter line revealed that pancreatic TH signaling is active in

islets during the larval-juvenile transition. Therefore, we next investigated how endogenous

TH signaling affects α cell and β cell development and glucose homeostasis during the larval-

juvenile transition. Towards this end, we used the NTR/Mtz system to ablate the thyroid in

tg:nVenus-2a-nfsb animals (22, 29). We treated tg:nVenus-2a-nfsB fish with Mtz at 20 dpf

and analyzed their phenotypes at 30 dpf (Fig. 6A). Overall eGFP expression of the TH

reporter was reduced, and TH signaling was not detected in the islets of Mtz treated animals

at 30 dpf (Fig. 6B). Having established that thyroid ablation eliminated TH signaling in

endocrine cells, we next analyzed its functional consequences. Islet morphology was

unaltered in thyroid-ablated zebrafish (Fig. 6C). Interestingly we found that compared to the

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unablated controls, the thyroid-ablated zebrafish exhibited elevated glucose levels (Fig. 6D),

that insulin expression levels were reduced (Fig. 6E), and that glucagon expression levels

were increased (Fig. 6E). Furthermore, expression levels of neurod1, pax6b, mnx1 and pax4

were reduced, and expression levels of arxa were increased in the digestive organs (liver,

pancreas and intestine) in thyroid-ablated zebrafish (Fig. 6E). These results suggest that

endogenous TH regulates glucose homeostasis during the larval-juvenile transition by

modulating the expression of endocrine genes including insulin and glucagon.

T3 regulates glucose metabolism by stimulating Insulin secretion and suppressing

Glucagon secretion in adults

Analysis of the TH reporter line revealed that pancreatic TH signaling is active in

adult islets. Therefore, we next investigated how TH signaling affects α cell and β cell

function and glucose metabolism in adults. 10 µl of a 10 nM T3 solution was injected

intraperitoneally (IP) daily for three consecutive days. After the third injection, the fish were

starved for 24 h and then challenged with an IP injection of 10 µl of a 2.5% glucose solution.

Blood glucose was measured after 45 mins (Fig. 7A). Under basal conditions as well as after

glucose challenge, T3 pretreatment reduced blood glucose levels (Fig. 7B). Next, pancreata

were dissected and the expression levels of insulin, glucagon a and glucagon b were analyzed

by qPCR. T3 upregulated expression of insulin, and downregulated expression of glucagon a

(Fig. 7C), consistent with the effects observed in larvae. T3 did not affect the expression

levels of glucagon b (Fig. 7C). To address whether altered gene expression levels led to

functional effects on hormone secretion, serum Insulin and Glucagon levels were quantified

by dot-blot. Serum Insulin levels were higher in animals treated with T3 compared to

controls (Fig. 7D and F). In contrast, serum Glucagon levels were reduced in fish treated

with T3 compared to controls (Fig. 7E and F). In addition, we investigated how TH signaling

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affects α cell and β cell function and glucose metabolism in adults by analyzing hypothyroid

animals. Mtz was injected intraperitoneally in tg:nVenus-2a-nfsb fish. Beginning two days

following the Mtz injection, the fish were starved for an additional 24 h and then challenged

with a 10 µl IP injection of 1% glucose (Fig. 7G). Under basal conditions as well as after

glucose challenge, blood glucose levels were elevated in hypothyroid zebrafish (Fig. 7H).

The expression levels of insulin, glucagon a and glucagon b in the pancreas of hypothyroid

zebrafish were analyzed by qPCR. The expression levels of insulin were reduced while the

expression levels of both glucagon a and glucagon b were significantly increased (Fig. 7I).

Furthermore, serum Insulin levels were reduced (Fig. 7J and L), and serum Glucagon levels

were elevated in hypothyroid zebrafish (Fig. 7K and L). Thus, TH mediated effects on

pancreatic endocrine hormone expression induce an anti-hyperglycemic islet secretion profile

that improves glucose tolerance.

Discussion

In this study, we investigated the role of TH signaling during pancreas development in

vivo. We found that TH signaling activates insulin expression in β cells and inhibits

glucagon expression in α cells. TH also increases the expression of a transcriptional network

including the pax6b and mnx1genes, whose protein products regulate insulin expression.

Pax6, a pan-endocrine factor, directly regulates β cell specific mnx1 expression through a

regulatory element in the mnx1 promoter (27). While mnx1 is expressed specifically in β

cells (27), pax6b is expressed in all endocrine cell types including β cells (25), and both

Mnx1 and Pax6b regulate insulin expression. Our data are consistent with a model whereby

TH induction of pax6b expression induces mnx1 expression to drive insulin expression in β

cells. Furthermore, TH increases the expression of pax4 and inhibits the expression of arxa,

whose protein products regulate glucagon expression. Arx and Pax4 are mutually

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antagonistic factors that modulate α cell development and glucagon expression (30). Our

data suggest that TH can tilt this balance in favor of Pax4, thus leading to a decrease in α cell

number and glucagon expression.

The TH/Pax6 pathway also appears to be at play in mammalian tissues. Exogenous

TH signaling can stimulate Insulin expression in db/db mice (31), and Pax6 expression is

reduced in TH deficient mice during fetal neurogenesis (32). Thus Pax6 might be one of the

common targets of TH signaling in pancreatic and neural tissues and might explain the

coordinated effects of TH signaling during embryogenesis, tissue differentiation, and organ

maturation. However, it remains to be investigated whether TH mediated Pax4 regulation is

also at play in mammalian tissues.

TH modulates the differentiation and function of various tissues including the

pancreas. However, it has been unclear when and in which cell types TH signals. Our data

using a novel TH responsive transgenic signaling reporter line reveal that in zebrafish, β cells

become TH responsive by 25 dpf and α cells by 30 dpf. These stages correspond to a critical

larval-juvenile transition, when zebrafish acquire an adult morphology and physiology (2, 3,

7, 33). This period is thought to be equivalent to the suckling to weaning transition in

mammals when various tissues undergo morphological and functional changes (2-7). During

this transition, rodent β cells acquire progressively higher glucose sensitivity and secretory

capacity (9). Our data indicate that islet endocrine cells are competent to respond to TH

during larval stages and that their maturation is stimulated by exposure to TH during and

after the larval-juvenile transition. Although TH signaling has been implicated in β cell

maturation (14), it is unclear to what extent endogenous TH signaling contributes to

mammalian pancreatic development and function. TH signaling is first detected in zebrafish

β cells during the larval-juvenile transition and remains active in adult islets. We found that

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TH signaling increases pax6 expression, which is required for maintaining the mature β cell

state in rodent islets (34-36). Thus, TH signaling may play a conserved role in the functional

maturation of β cells as well as the maintenance of β cell function in adult islets. In mammals,

Pax6 promotes and maintains β cell maturation by inducing a transcriptional network that

includes MAFA (34-37). However, in zebrafish we did not detect mafa expression in

pancreatic β cells (Supplemental Figure 4). Therefore, in zebrafish, Pax6b may use MafA-

independent pathways to induce β cell maturation. A complete understanding of the diversity

of TH-dependent regulatory mechanisms in vertebrates may reveal novel therapeutic

strategies to promote β cell maturation.

Interestingly, our data indicate that islet endocrine cells are competent to respond to

exogenous T3 during larval stages, but that endogenous T3 levels are not sufficient to

stimulate islet cells before 25 dpf. In addition, during development, β and α cells become TH

responsive at different stages (β cells by 25 dpf and α cells by 30 dpf), despite the fact that

both cell types have the competence to sense and respond to exogenous T3. Thus, islet cells

may have different thresholds for endogenous TH signaling. TH is synthesized in the

thyroid as thyroxine (T4), an inactive form for THR activation. Next, T4 is converted into T3,

a potent THR activator, by type 1 and type 2 deiodinases (D1 and D2, respectively) in

peripheral target tissues, and finally converted into the inactive T2 form by a type 3

deiodinase (D3) (38). In rat islets, D1 protein levels are much lower at postnatal day 7

compared to adult (14). In contrast, D3 shows a reciprocal pattern (14). Thus, expression of

key TH metabolic enzymes may explain the tissue specificity of the TH response.

Furthermore, in zebrafish, TH production dramatically increases during the larval-juvenile

transition (39). Thus, elevation of TH production/release and elevation of T3 by changes in

TH metabolism in target peripheral tissues may produce unique TH responses in each

tissue/cell type during the larval-juvenile transition. In addition, it has been reported that

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Thrα and Thrβ have different expression patterns in rat islets (14). Thrβ is also regulated by

TH and marks mature tissues. Thus, understanding the differences between THRA and B

may provide additional information necessary to understand the molecular mechanisms of

tissue specific transcriptional regulation and tissue maturation by TH.

Neonatal β cells differ from adult β cells in the expression of key metabolic genes

including those involved in mitochondrial ATP production (11). TH has been shown to

stimulate mitochondrial biogenesis and function through activation of PGC-1 and NRF1

expression (40). TH-stimulated rat islets show an increase in pgc1a expression at P7

although it is unclear which cell types are TH responsive (14). Thus, TH signaling may

increase Insulin secretion in β cells through stimulation of mitochondrial function.

Mitochondrial ATP production is also implicated in the inhibition of Glucagon secretion by α

cells (41). Thus, increased mitochondrial metabolic activity in TH-responsive islet cells

could explain the hyperinsulinemia and hypoglucagonemia that we observed in TH-treated

adults. Further analysis of the link between TH signaling and mitochondrial function may

provide key insights into the regulation of islet maturation and function.

TH induced stimulation of Insulin secretion and inhibition of Glucagon secretion led

to reduced fasting glucose levels and increased glucose tolerance in larvae and adults.

Unravelling the regulation of TH signaling will help establish its role in the pathophysiology

of diseases including diabetes. In human, both hypothyroidism and hyperthyroidism increase

the incidence of diabetes (42). However, preclinical studies indicate that TH therapy

attenuates hyperglycemia and Insulin resistance in the db/db diabetic mouse model (31).

Therefore, exogenous TH administration within the physiological range may have therapeutic

applications. The acquisition of TH responsiveness during the functional maturation of islets

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during the larval-juvenile transition provides an in vivo platform to develop

antihyperglycemic therapies modulating this pathway.

Acknowledgement

This work was supported by funds from the Max Planck and EU society to D.Y.R.S.

No potential conflicts of interest relevant to this article were reported.

H.M. and D.Y.R.S. conceived the project. H.M. S.T.M, D. H. and D.Y.R.S.

contributed to discussion of data and manuscript preparation. H.M., S.T.M., and Y.Z

generated data. H.M. and D.Y.R.S. are the guarantors of this work and, as such, had full

access to all the data presented in the study and take responsibility for the data and the

accuracy of the data analysis.

We thank Dirk Meyer (University of Innsbruck) for the mnx1:eGFP line, David

Parichy (University of Washington) for the tg:nVenus-2a-nfsB line, Yu Hsuan Carol Yang,

Michelle Collins, Jasmin Gäbges and Einat Blitz (MPI-BN) for critical reading of the

manuscript, Sabine Fischer for the animal protocols, all members of the Stainier lab for

helpful discussion and the fish facility staff for fish care.

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Figure legends

Figure 1. T3 stimulates insulin expression, reduces glucose levels, and improves glucose

tolerance. (A-C) Zebrafish were treated with 0, 0.1, 1, 10 or 100 nM T3 from 4 to 7 dpf and

analyzed at 7 dpf for insulin promoter activity (B) and for glucose levels (C). Note that

insulin promoter activity was significantly enhanced by 10 nM T3 treatments, and that

glucose levels were reduced by 10 and 100 nM T3 treatments. (D-F) Zebrafish were treated

with 10 nM T3 from 4 dpf and analyzed for insulin promoter activity (E) and glucose levels

(F) at 5, 6, 7 and 8 dpf. Note that insulin promoter activity was enhanced at 7 and 8 dpf by

T3 treatment (E), and that glucose levels were reduced at 6, 7 and 8 dpf by T3 treatment (F).

(G and H) Larvae were first treated with 10 nM T3 from 4 to 7 dpf, then with 5 % glucose for

1 hour and finally analyzed for glucose levels. Glucose tolerance was compared after

treatment with and without T3 (H). Values of bioluminescence signal were derived from 5

wells per condition, with 3 animals per well. Values of glucose were derived from 5 wells

per condition, with extracts from 20 animals per well. *, P<0.05; **, P<0.01 compared to

controls by Turkey-Kramer HSD test after ANOVA; ns, not significant. AU, arbitrary units.

Figure 2. T3 enhances insulin expression levels, but does not change insulin expression

pattern or β cell numbers. (A) Expression of insulin (ins) mRNA was compared between

control and T3-treated larvae by qPCR. Endogenous ins expression was enhanced after 3

days of T3 treatment (7 dpf). (B and C) Expression pattern of ins was compared between

control (B) and T3-treated (C) larvae at 7 dpf by in situ hybridization. ins expression was

detected only in pancreatic β cells (arrow), but not in other tissues in control (B) and T3-

treated (C) larvae. (D-H) Tg(ins:eGFP) animals were treated with 10 nM T3 starting at 4 dpf

and analyzed at 7 dpf. eGFP expression mimicked the endogenous ins expression pattern in

control (D) and T3-treated (E) larvae. However, eGFP intensity was enhanced after T3

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treatment (D-H). (I-K) Tg(ins:H2BGFP) animals were treated with 10 nM T3 starting at 4

dpf and analyzed at 7 dpf. Note that the β cell number did not change with T3 treatment (K).

Scale bars in B, C, D and E, 200 µm; scale bars in F,G, I and J, 10 µm. **, P<0.01 compared

to controls by Turkey-Kramer HSD test after two-way ANOVA in A. *, P<0.05 compared to

controls by Student’s t-test in H and K. ns, not significant.

Figure 3. T3 represses glucagon expression. (A) Expression of glucagon a (gcga) and

glucagon b (gcgb) mRNA was compared after treatment with and without T3 by qPCR. The

expression of gcga and gcgb was reduced at 5, 6 and 7 dpf by T3 treatment. (B) Expression

patterns of gcga and gcgb were compared after treatment with and without T3 by in situ

hybridization. Both gcga and gcgb expression levels appeared to be reduced after T3

treatment. (C and D) Tg(gcga:eGFP) animals were treated with 10 nM T3 starting at 4 dpf

and the eGFP expression levels in the islets were analyzed at 7 dpf. Note that eGFP intensity

was decreased after T3 treatment (D). (E) Tg(gcga:eGFP) positive cell number was

compared after treatment with and without T3. Note that the α cell number was decreased

after T3 treatment. Scale bar in B, 100 µm; scale bar in C, 10 µm. **, P<0.01 compared to

controls by Turkey-Kramer HSD test after two-way ANOVA in A. *, P<0.05 compared to

control by Student’s t-test in D and E. ns, not significant.

Figure 4. T3 modulates the expression of endocrine differentiation markers. (A-F)

Tg(pax6b:eGFP) (A and B), Tg(neurod1:eGFP) (C and D) and Tg(mnx1:eGFP) (E and F)

animals were treated with T3 to investigate its effect on three genes (pax6b, neurod1 and

mnx1) involved in endocrine cell differentiation. Note that eGFP intensities were enhanced

in Tg(pax6b:eGFP) and in Tg(mnx1:eGFP) animals, but not in Tg(neurod1:eGFP) animals

after T3 treatment. (G) qPCR analysis showed that T3 treatment enhances the expression of

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pax6b, mnx1, and pax4 and reduces the expression of arxa at 6 dpf. Scale bars, 10 µm. *,

P<0.05; **, P<0.01 compared to controls by Student’s t-test. ns, not significant.

Figure 5. Pancreatic α and β cells are targets of thyroid hormone signaling during and

after larval development. (A) Schematic diagram of thyroid hormone reporter construct.

(B-E) Thyroid hormone reporter expression was compared with Glucagon and Insulin

expression in islets at 20 (B), 25 (C) and 30 (D) dpf and in adults (E) by

immunohistochemistry. Note that strong eGFP signal could be detected in both Glucagon

(red arrows) and Insulin (white arrows) positive cells at 30 dpf and in adults. Weak eGFP

signal in Insulin positive cells, but not in Glucagon positive cells, was also present at 25 dpf

(white arrows). (G) Ratio of TRE:eGFP positive cells to Glucagon positive cells as well as

Insulin positive cells. Values were derived from 5 fishes per developmental time point, with a

minimum of 3 sections per fish. Scale bars, 5 µm. *, P<0.05; **, P<0.01 compared to 20 dpf

by Turkey-Kramer HSD test after ANOVA.

Figure 6. Endogenous thyroid hormone regulates endocrine marker expression and

glucose levels during the larval to juvenile transition. (A) Schematic detailing the time

course of Mtz treatment in Tg(tg:nVenus-2a-nfsB) line to generate hypothyroid zebrafish.

(B) Thyroid hormone reporter expression was reduced in 30 dpf Tg(tg:nVenus-2a-

nfsB)/Tg(6xTRE-bglob1:eGFP) fish after Mtz treatment. (C) Morphology of islets in

hypothyroid zebrafish at 30 dpf. No morphological differences could be detected in the

pancreatic islets between control and Mtz-treated animals. (D) Free glucose level

measurements at 30 dpf show that hypothyroid zebrafish exhibited elevated glucose levels.

(E) qPCR analysis of 30 dpf animals treated with or without Mtz reveals that thyroid ablation

resulted in a marked decrease of ins, neurod1, pax6b, mnx1, and pax4 expression, while the

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expression of gcga and arxa was significantly increased. Triangles indicate principal islet.

Arrows point to secondary islets. Scale bars, 200 µm. *, P<0.05; **, P<0.01 compared to

controls by Student’s t-test. ns, not significant.

Figure 7. Thyroid hormone signaling regulates glucose homeostasis by stimulating

Insulin secretion and repressing Glucagon secretion in adult zebrafish. (A) Schematic

time course of T3 and glucose injections. (B) Blood glucose levels in control and T3 treated

adult fish before and after glucose injections. (C) insulin and glucagon expression in adult

pancreas by qPCR after T3 treatment. (D) Plasma Insulin levels after T3 treatment compared

to control. (E) Plasma Glucagon levels after T3 treatment compared to control. (F)

Quantification of D and E. (G) Schematic time course of Mtz and glucose injections. (H)

Blood glucose levels in Tg(tg:nVenus-2a-nfsB) animals treated with Mtz before and after

glucose injection. (I) insulin and glucagon expression in adult pancreas by qPCR in

Tg(tg:nVenus-2a-nfsB) animals treated with Mtz. (J) Plasma Insulin levels in Tg(tg:nVenus-

2a-nfsB) animals treated with Mtz. (K) Plasma Glucagon levels in Tg(tg:nVenus-2a-nfsB)

animals treated with Mtz. (L) Quantification of J and K. *, P<0.05; **, P<0.01 compared to

controls by Student’s t-test. ns, not significant.

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Supplemental Figure 1. T3 regulates glucagon expression in an Insulin-independent

manner. Expression levels of gcga and gcgb mRNA were assessed in 6 dpf insulin mutant

larvae after or without T3 treatment. The expression levels of gcga and gcgb were reduced

after 2 days of T3 treatment.

Supplemental Figure 2. Generation of thyroid hormone reporter zebrafish. (A)

Schematic diagram of thyroid hormone reporter construct. Four different minimal promoters

(cfos, bglob1, tk and Pmini) were tested as candidates to build the optimal reporter construct.

(B) Time course of transient injection assays to compare reporter activities in zebrafish

embryos. (C and D) Results of transient injection assays. Note that the construct with the

bglob1 minimal promoter showed the highest reporter activation upon T3 treatment. (E)

Schematic diagrams of thyroid hormone reporter constructs. (F) Results of transient injection

assays. Note that the 6xTRE-bglob1:eGFP construct was used to generate the thyroid

hormone reporter zebrafish line.

Supplemental Figure 3. T3 induces ubiquitous eGFP expression in Tg(6xTRE-

bglob1:eGFP) larvae. (A-E) Tg(6xTRE-bglob1:eGFP) animals were treated with T3 starting

at 4 dpf and eGFP expression analyzed at 7 dpf. Note that eGFP was strongly induced by T3

treatment. C and E show higher magnification images after dissecting out the digestive

organs. p, pancreas; l, liver; i, intestine. Dotted lines in C outline the pancreas. Scale bars,

200 µm.

Supplemental Figure 4. mafa expression in adult zebrafish pancreas. Two color in situ

hybridization was performed with anti-insulin and anti-mafa probes. We could not detect any

overlapping expression of mafa (blue) and insulin (brown), indicating that in zebrafish mafa

is not expressed in adult beta cells, but only in the exocrine pancreas.

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Supplemental Material

Luciferase assay

After T3 treatment, samples were incubated in Steady Glo (Promega) as described

previously (1). Bioluminescence signal was analyzed by FLUOstar Omega (BMG

LABTECH).

Glucose measurements and treatments

After T3 treatment, 20 larvae or 10 juveniles per group were ground up and free

glucose levels were determined by using a glucose assay kit (BioVision) as described

previously (1). For the glucose tolerance tests, samples were incubated in 5% glucose in egg

water for 1 hr and then free glucose was measured using the glucose assay kit. For the graphs

shown in Figure 1 and Figure 6D, each sample was normalized to a control sample. For

blood glucose measurements, adult zebrafish were anesthetized in tricaine and then cut by the

anal fin with a knife. Whole blood was analyzed immediately with a glucometer (Abbott).

Glucose treatment of adult zebrafish was performed by injecting 10 µl of 1 or 2.5 % glucose

intraperitoneally.

qRT-PCR

Whole animals were used for RNA isolation from larval zebrafish. For RNA

isolation from juvenile zebrafish, the gut, liver and pancreas were dissected out. For RNA

isolation form adult zebrafish, the pancreases were dissected out. Total RNA from control

and T3-treated larvae was extracted by TRIZOL reagent (Ambion) and then cleaned with

RNA Clean & Concentrator-5 (ZYMO RESEARCH) to remove DNA contamination.

Reverse transcription/polymerase chain reaction (RT–PCR) was performed by using

SuperScript II First-Strand Synthesis System (Invitrogen) according to the manufacturer’s

instructions with 500 ng total RNA as a template.

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qRT-PCR with 2xDynamo Color Flash SYBR Green master mix (Thermo Scientific)

was carried out to quantify gene expression levels on a CFX connect Real-time System (Bio

Rad) with gene-specific primers (Supplemental Table 1). Each sample was normalized to a

control gene (actb). Cq values are listed in Supplemental Table 2.

In situ hybridization

Partial cDNA’s encoding zebrafish insulin, glucagon a and glucagon b were obtained

by reverse transcription (RT)-PCR with RNA extracted from zebrafish at 3 days post

fertilization (dpf) with gene-specific primers (Supplemental Table 1). insulin PCR products

were cloned into pCRII-TOPO vector (Invitrogen) and verified by sequencing. glucagon a

and glucagon b PCR products were cloned into pGEM-T-easy vector (Promega) and verified

by sequencing. To synthesize antisense RNA probes, these plasmids were linearized with

BamHI for insulin, SacII for glucagon a, and SpeI for glucagon b and transcribed with T7

RNA polymerase for insulin and glucagon b and Sp6 RNA polymerase for glucagon a

(Roche Applied Science). Whole-mount in situ hybridizations were performed as described

previously (2).

Fluorescence imaging and immunohistochemistry

Fluorescence images were acquired with a LSM700 or LSM780 laser scanning

confocal microscope (Carl Zeiss), and fluorescence intensity for each transgenic line was

calculated using the ZEN software. Cell numbers were counted manually for ins:H2BGFP or

using the Imaris software (Bitplane) for gcga:eGFP.

Immunohistochemistry was performed as described previously (3). The following

primary antibodies were utilized at the indicated dilutions in 1% FBS in PBS-containing

0.1% Triton X-100 (PBTx): guinea pig anti-Insulin (Thermo Scientific) at 1:300, monoclonal

mouse anti-Glucagon (Sigma Aldrich) at 1:100 and chicken anti-eGFP (Thermo Scientific) at

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1:500. AlexaFluor-conjugated secondary antibodies (Invitrogen) were used at 1:500-1:1000

dilution with 1% FBS in PBTx.

Plasmid construction, Transient injection assays and transgenesis

To generate Tg(ins:Luc2;cryaa:mCherry)gi3

, luc2 was placed under the control of a

1.1 kb insulin promoter element. The -0.5cryaa:mcherry transgenesis marker was inserted

into the vector in the reverse orientation.

For the thyroid hormone responsive element, we utilized the promoter from the

Xenopus laevis TH/bzip gene, which contains two THR response elements (4). To produce

the reporter plasmid 6xTHbzipTREs-cFos:eGFP, TREs-cFos element containing 3 copies of

TH/bzip promoter (6 TREs) and a mouse cfos minimal promoter was amplified and inserted

into the cFos:eGFP vector (5). Each 6xTHbzipTREs-bglob1:eGFP, 6xTHbzipTREs-tk:eGFP,

and 6xTHbzipTREs-Pmini:eGFP was generated by replacing the cFos promoter of

6xTHbzipTREs-cFos:eGFP with a rabbit bglob1 minimal promoter (6), and a HSV thymidine

kinase minimal promoter (tk) minimal promoter (7) and synthetic mini promoter (Pmini)

promoter (8). bglob1:eGFP was generated by replacing the 6xTHbzipTREs-Pmini region of

6xTHbzipTREs-Pmini:eGFP with a bglob1 minimal promoter. To produce 2xTHbzipTREs-

bglob1:eGFP, a copy of the TH/bzip promoter was replaced by the 6xTHbzipTREs part of

6xTHbzipTREs-bglob1:eGFP. To produce 6xTHbzipTREs (mut)-bglob1:eGFP, a mutated

TH/bzip TREs -bglob1 element, containing 3 copies of a mutated TH/bzip promoter (4) and a

bglob1 promoter was amplified and placed instead of the 6xTHbzipTREs-cFos region of the

6xTHbzipTREs-cFos:eGFP construct.

One-cell stage zebrafish embryos were microinjected with each promoter construct

using a micromanipulator. The injected embryos were transferred to culture dishes and

reared in egg water at 28 °C. Injected embryos were treated with T3 starting at 24 hpf and

analyzed for eGFP signal at 60 hpf.

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To generate Tg(ins:Luc2;cryaa:mCherry)gi3

, the ins:Luc2;cryaa:mCherry plasmid

was co-injected with I-SceI meganuclease into one-cell stage embryos as described

previously (9). To generate Tg(6xTRE-bglob1:eGFP)bns91Tg

, the 6xTHbzipTREs-

bglob1:eGFP plasmid was co-injected with tol2 mRNA into one-cell stage embryos as

described previously (10). Multiple stable lines were established. A single representative

line was used for all experiments.

Plasma Insulin and Glucagon detection

After making a cut at the level of the anal fin with a knife, 1 µl of blood was taken per

fish and placed into a tube containing 5 µl heparin. The blood of 10 fish per condition was

pooled and centrifuged at 13700 g at 4 ºC for 15 mins to separate the cells and plasma. An

equal volume of plasma sample was used for dot blotting. Dot blots were performed as

described previously (11). For Insulin and Glucagon detection, guinea pig anti-Insulin

(1:200) (Thermo Scientific) and monoclonal mouse anti-Glucagon (1:600) (Sigma-Aldrich)

antibodies were used. The membrane was analyzed with a ChemiDoc MP imaging system

(Bio Rad).

Supplemental references

1. Gut P, Baeza-Raja B, Andersson O, Hasenkamp L, Hsiao J, Hesselson D, Akassoglou K,

Verdin E, Hirschey MD, Stainier DY. Whole-organism screening for gluconeogenesis

identifies activators of fasting metabolism. Nat Chem Biol 2013;9: 97–104.

2. Cerveny K, Houart C. Wholemount in situ Hybridization and Antibody Staining. Neural

Development and Genetics of the Zebrafish 2009:103-107.

3. Matsuda H, Parsons MJ, Leach SD. Aldh1-expressing endocrine progenitor cells regulate

secondary islet formation in larval zebrafish pancreas. PLoS One 2013;8:e74350.

of 48 Diabetes

6

4. Furlow JD, Brown DD. In vitro and in vivo analysis of the regulation of a transcription

factor gene by thyroid hormone during Xenopus laevis metamorphosis. Mol Endocrinol

1999;13:2076-2089.

5. Fisher S, Grice EA , Vinton RM , Bessling SL , Urasaki A , Kawakami K , McCallion

AS. Evaluating the biological relevance of putative enhancers using Tol2 transposon-

mediated transgenesis in zebrafish. Nat Protoc 2006;1:1297–1305.

6. Parsons MJ, Pisharath H, Yusuff S, Moore JC, Siekmann AF, Lawson N, Leach SD.

Notch-responsive cells initiate the secondary transition in larval zebrafish pancreas. Mech

Dev 2009;126:898-912

7. Shimizu N, Kawakami K, Ishitani T. Visualization and exploration of Tcf/Lef function

using a highly responsive Wnt/β-catenin signaling-reporter transgenic zebrafish. Dev Biol

2012;370:71-85.

8. Weger BD, Weger M, Nusser M, Brenner-Weiss G, Dickmies T. A chemical screening

system for glucocorticoid stress hormone signaling in an intact vertebrate. ACS Chem

Biol 2012;7:1178-1183.

9. Thermes V, Grabher C, Ristoratore F, Bourrat F, Choulika A, Wittbrodt J, Joly JS. I-SceI

meganuclease mediates highly efficient transgenesis in fish. Mech Dev 2002;118:91-98.

10. Kawakami K. Transgenesis and gene trap methods in zebrafish by using the Tol2

transposable element. Methods Cell Biol 2004;77:201–224.

11. Olsen AS, Sarras MP Jr, Intine RV. Limb regeneration is impaired in an adult zebrafish

model of diabetes mellitus. Wound Repair Regen 2010;18:532-542.

12. Hesselson D, Anderson RM, Beinat M, Stainier DY. Distinct populations of quiescent

and proliferative pancreatic beta-cell identified HOTcre mediated labeling. Proc Natl

Acad Sci U S A 2009;106:14896-14901.

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13. Ye L, Robertson MA, Hesselson D, Stainier DY, Anderson RM. Glucagon is essential

for alpha cell transdifferentiation and beta cell neogenesis. Development 2015;142:1407-

1417.

14. McCurley AT, Callard GV. Characterization of housekeeping genes in zebrafish: male-

female differences and effects of tissue type, developmental stage and chemical treatment.

BMC Mol Biol 2008;9: 102.

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gcga

gcgb

0

1

mR

NA

leve

ls

controlT3

Matsudaetal.,SupplementalFigure1

** *

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6xTHbzipTREs

cfosbglob1tk

PminieGFPxxxminiP

A

B24 60hpf0

+T3

+T3

-T3

+T3

cfos

bglob1

0102030405060708090100

cfos

bglob1 tk

Pmininu

mbe

rofe

GFP+

embryos/total

embryos(%)

withoutT3

withT3

eGFPbglob1bglob1:eGFP

2xTRE-bglob1:eGFP eGFPbglob1

eGFPbglob16xTRE-bglob1:eGFP

E

eGFPbglob16xTRE(mut)-bglob1:eGFP

-T3

D

0102030405060708090100

noTRE

2xTR

E6xTR

E6xTR

E(m

ut)

numbe

rofe

GFP+

em

bryos/totalembryos(%

)

withoutT3

withT3

F

C


cfos

bglob1 tk

Pmini

of 48 Diabetes

4 7dpf0

+T3

control

T3

. . . ..... . . .......

......

. . .

p

li

A

B C

D E


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insmafa


of 48 Diabetes

Supplemental Table 1. primers sequence

(A) Primers for qPCR

primer name sequence (5'-->3') reference

insulin -f TTTAAATGCAAAGTCAGCCACCTCAG 12

insulin -r GGCTTCTTCTACAACCCCAAGAGAGA 12

gcga -f AAGGCGACAGCACAAGCACA 13

gcga -r GCCCTCTGCATGACGTTTGACA 13

gcgb -f GGAGACCAGGAGAGCACAAG

gcgb -r TGCAGGTA CGAGCTGACATC

neurod1 -f CTTTCAACACACCCTAGAGTTCCG

neurod1 -r GCATCATGCTTTCCTCGCTGTATG

pax6b -f GCCAGGACAACCAAATCAAGACG

pax6b -r GTGAAGGACGTTCTGTTTCTCTGC

mnx1 -f GAAGAGGAGCGGTGACACTC

mnx1 -r GGTCTGCACTTTTGGTGGAT

pax4 -f GAGAGAGCCTGCAGAGGAGA

pax4 -r TGTTGCTGAAAATGGCTCTG

arxa-f GGAGAGACTGCTGGACCAAG

arxa-r TTCATCACTCTGGCTGAACG

gapdh -f GTGGAGTCTACTGGTGTCTTC 14

gapdh -r GTGCAGGAGGCATTGCTTACA 14

(B) Primers for ISH probe

primer name sequence

insulin -f CCATATCCACCATTCCTCGCC

insulin -r CAAACGGAGAGCATTAAGG CC

gcga -f ATTTGCTGGTGTGTGTTGGA

gcga -r TAGTG TTTGGCACAGGGTGA

gcgb -f GAGAAAACCAGCGAGACGAC

gcgb -r GCATGATTCATACAGCACTGG

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larval

Cq

5 dpf

actb 25.06

25.11

actb+ 24.99

24.88

ins 25.76

25.73

ins+ 25.9

25.66

6 dpf

actb 24.63

24.69

actb+ 24.53

24.6

ins 25.62

25.63

ins+ 25.77

25.58

7 dpf

actb 24.78

24.85

actb+ 24.96

24.93

ins 25.48

25.23

ins+ 24.2

23.92

5 dpf

actb 25.06

25.11

actb+ 24.99

24.88

gcga 26.42

26.67

gcga+ 27.65

27.42

gcgb 28

27.89

gcgb+ 28.39

28.5

6 dpf

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actb 24.63

24.69

actb+ 24.53

24.6

gcga 26.01

25.82

gcga+ 26.4

26.71

gcgb 28.26

28.16

gcgb+ 28.56

28.52

7 dpf

actb 24.78

24.85

actb+ 24.96

24.93

gcga 28.52

28.87

gcga+ 28.81

28.92

gcgb 29.63

29.61

gcgb+ 29.89

30.08

6 dpf

actb 28.5

28.49

actb+ 28.72

28.26

neurod1 29.21

29.25

neurod1+ 29.03

28.75

pax6b 28.87

28.61

pax6b+ 27.75

27.82

mnx1 30.84

31.25

mnx1+ 29.81

30.02

pax4 29.32

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29.24

pax4+ 28.11

28.27

arxa 29.53

30.12

arxa+ 31.02

31.23

juvenile

30 dpf

actb 24.46

24.11

24.16

actb+ 24.74

24.39

24.3

ins 28.08

28.03

28.02

ins+ 29.13

28.72

28.71

gcga 34.85

35.07

34.9

gcga+ 33.64

33.19

33.26

gcgb 28.28

28.51

28.48

gcgb+ 28.06

28.31

28.17

neurod1 34.54

34.69

34.7

neurod1+ 35.27

35.62

35.37

pax6b 33

32.49

32.7

pax6b+ 34.88

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34.78

34.32

mnx1 33.26

33.25

32.75

mnx1+ 34.61

34.14

34.1

pax4 33.07

32.42

32.43

pax4+ 34.56

34.38

33.64

arxa 35.82

35.06

35.41

arxa+ 34.22

34.5

34.89

adult

actb 25.83

25.88

actb+ 25.75

25.79

ins 24

23.91

ins+ 23.15

22.73

gcga 23.16

23.25

gcga+ 24.23

24.3

gcgb 26.18

26.7

gcgb+ 26.15

26.22

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page 1 of 48 diabetes...1 thyroid hormone coordinates pancreatic islet maturation during the...

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