TIGAR (Tp53-induced glycolysis and apoptosis regulator) was originally identified as a p53-inducible protein that functions as a fructose-2,6-bisphosphatase (F2,6P) but subsequently shown to have phosphatase activities for a variety of phosphorylated metabolic intermediates and allosteric regulators including 2,3-bisphospholgycerate (2,3BPG), 2-phosphoglycerate, phosphoglycolate, and phosphoenolpyruvate (Bensaad et al., 2006; Rigden, 2008; Bolaños, 2014; Tang et al., 2021). Due to its F2,6P dephosphorylation activity, TIGAR is generally considered a suppressor of glycolysis as F2,6P is a potent allosteric activator for 6-phosphofructo-1-kinase (PFK1), the rate-limiting step in glycolysis. However, the dephosphorylation of 2,3BPG to generate 3-phosphoglycerate would also be expected to increase glycolysis (Bolaños, 2014). In addition, the role of TIGAR in regulating carbohydrate metabolism is also complicated by the presence of the related phosphofructokinase bis-phosphatase (PFKBP) family that has both F6P 2-kinase and F2,6P bisphosphatase activities (Mor et al., 2011). Thus, the biological readout of TIGAR function is likely to be highly cell context dependent. In this regard, TIGAR has been reported to differentially modulate multiple different pathophysiological outcomes. For example, in the brain TIGAR expression was reported to protect against ischemic/reperfusion injury (Li et al., 2021), whereas in the heart TIGAR deficiency protected myocardial infarction (Hoshino et al., 2012) and in pressure-overloaded hearts (Okawa et al., 2019). In cancer models, TIGAR plays a complex role in the formation and progression of different types of cancer via suppressing aerobic glycolysis and controlling reactive oxygen species (ROS) production (Tang et al., 2021). Recently, it has been reported that TIGAR can both enhance the development of premalignances and suppress the metastasis of cancer invasion by the way of inhibition of ROS production (Cheung et al., 2020).

As carbohydrate metabolism and glycolysis are essential normal physiological processes in all cells and tissues, we have examined the metabolic phenotype of whole-body and tissue-specific TIGAR knockout mice. Surprisingly, we have observed that TIGAR deficiency in cholinergic neurons of mice results in a marked protection against hypothermia following an acute cold challenge. At a molecular level, this results from enhanced acetylcholine levels and increased cholinergic signaling at the neuromuscular junction (NMJ) driving skeletal muscle shivering-induced thermogenesis.

Previously we reported that the TIGAR can modulate NF-kB signaling through a direct binding interaction and inhibition of the E3 ligase activity of the linear ubiquitin-binding assembly complex, LUBAC, in cultured cells (Tang et al., 2018). To examine the role of TIGAR in vivo, we initially generated whole-body Tigar knockout (TKO) mice. Surprisingly, however, during our phenotypic characterization we observed that the TKO male and female mice both display a remarkable resistance to cold induced-hypothermia. As shown in Figure 1, both male (panel A) and female (panel B) control mice when shifted from room temperature to a 4°C environment display the typical 4–5°C decline in core body temperature during the 1 hr acute exposure period. In contrast, the TKO male and female mice only display less than a 1°C decline in core body temperature under identical conditions. The level of Ucp1 mRNA (Figure 1—figure supplement 1A) and protein (Figure 1—figure supplement 1B) was not significantly different in interscapular brown adipose tissue (iBAT) at room temperature. Ucp1 mRNA did increase approximately 1.5-fold following 1 hr cold exposure, but this was not significantly different between the control and TKO mice. Although a very small amount of Ucp1 mRNA was detected in inguinal adipose tissue (iWAT) of control mice at room temperature, it was lower in the TKO mice (Figure 1—figure supplement 1C). Moreover, following 4°C exposure for 2 hr there was an increase in Ucp1 mRNA indicative of beige adipocyte induction. However, the induction of Ucp1 mRNA was significantly reduced in the TKO mice compared to the control mice, probably due to decreased sensitivity of cold-induced sympathetic activation (see Figure 7).

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Fresh gastrocnemius muscle (Gastr), epididymal adipose tissue (EWAT), inguinal white adipose tissue, and interscapular brown adipose tissue from Tigar^fl/fl and Tigar^fl/fl>/Adipoq^Cre mice were collected, and 30 μg of the tissue lysate were used for TIGAR (30 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section.

The raw image was used for Figure 1C. A detailed description of the raw images is shown in Source data 1.

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Figure 1—source data 2

Interscapular brown adipose tissues were collected from wild type (WT), whole-body Tigar knockout (TKO), Ucp1 knockout (UKO), and Ucp1 and Tigar double knockout (UTKO) mice, and 30 μg of the tissue lysate were used for TIGAR (30 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section.

The raw image was used for Figure 1E. A detailed description of the raw images is shown in Source data 1.

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Figure 1—source data 3

Interscapular brown adipose tissues were collected from wild type (WT), whole-body Tigar knockout (TKO), Ucp1 knockout (UKO), and Ucp1 and Tigar double knockout (UTKO) mice, and 30 μg of the tissue lysate were used for UCP1 (33 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section.

The raw image was used for Figure 1E. A detailed description of the raw images is shown in Source data 1.

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Figure 1—source data 4

Interscapular brown adipose tissues were collected from wild type (WT), whole-body Tigar knockout (TKO), Ucp1 knockout (UKO), and Ucp1 and Tigar double knockout (UTKO) mice, and 30 μg of the tissue lysate were used for vinculin (117 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section.

The raw image was used for Figure 1E. A detailed description of the raw images is shown in Source data 1.

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Figure 1—source data 5

Fresh heart, gastrocnemius muscle (Gastr), extensor digitorum longus (EDL muscle), and soleus muscle were collected from Myl1^Cre and Myl1^Cre/Tigar^fl/fl mice, and 30 μg of the tissue lysate were used for TIGAR (30 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section.

The raw image was used for Figure 1G. A detailed description of the raw images is shown in Source data 1.

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Figure 1—source data 6

Fresh heart, gastrocnemius muscle (Gastr), extensor digitorum longus (EDL muscle), and soleus muscle were collected from Myl1^Cre and Myl1^Cre/Tigar^fl/fl mice, and 30 μg of the tissue lysate were used for vinculin (117 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section.

The raw image was used for Figure 1G. A detailed description of the raw images is shown in Source data 1.

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Changes in tissue distribution of muscle and fat mass as well as modulation of basal energy expenditure have important roles in the control of thermogenesis. However, there was no significant difference in the relative fat mass or lean mass (Figure 1—figure supplement 1D) between the control and TKO mice. Hematoxylin and eosin (H&E) staining of oxidative soleus and glycolytic extensor digitorum longus (EDL) (Figure 1—figure supplement 1E) skeletal muscle also indicated the absence of any gross morphological differences between the control and TKO mice. Similarly, food intake (Figure 1—figure supplement 1F), spontaneous locomotor activity in the AMB + Z dimensions (Figure 1—figure supplement 1G), and energy expenditure (Figure 1—figure supplement 1H) were essentially identical.

As the TKO mice are whole-body knockouts, the temperature phenotype could result from an effect on a single or combinatorial number of cell types. We first generated Tigar^fl/fl mice that were crossed with Adipoq^Cre mice to generate adipocyte-specific TKO mice (Figure 1C). These mice had the identical temperature sensitivity as the control Tigar^fl/fl mice both displaying an approximately 4°C decline in core body temperature following exposure to 4°C for 1 hr (Figure 1D). To further rule out BAT as a contributor to the enhanced cold protection in the TKO mice, we crossed the cold-sensitive Ucp1-deficient mice (Enerbäck et al., 1997) with the TKO mice (both strains with C57BL/6J background) to produce wild type (WT), TKO, UKO, and Ucp1^-/-/Tigar^-/- double knockout (UTKO) mice (Figure 1E). As expected, UKO mice were highly cold sensitive compared to their WT littermates following a 1 hr acute cold exposure (Figure 1F). Interestingly, the UTKO mice displayed a strong cold resistance, which was not significantly different from the TKO littermates. These data strongly suggest that thermogenic adipose tissue is not responsible for the resistance to hypothermia in the TKO mice.

Since skeletal muscle plays a major role in heat production, we next generated skeletal muscle-specific TKO mice by crossing the Tigar^fl/fl mice with the skeletal muscle-specific myosin light polypeptide 1-Cre (Myl1^Cre) mice (Figure 1G). TIGAR is highly expressed in skeletal muscle, with the highest protein levels in depots containing glycolytic fibers (white gastrocnemius and EDL) skeletal muscle. The Myl1^Cre-driven TIGAR deletion resulted in efficient loss of TIGAR protein in all skeletal muscles examined with no effect on the heart (Figure 1G). Similar to the adipocyte-specific TKO mice, skeletal muscle Tigar-deficient mice had no significant resistance to acute cold exposure (Figure 1H). Moreover, we did not find any significant change in skeletal muscle mRNA expression of sarco-/endoplasmic reticulum Ca²⁺-ATPase (SERCA1, Atp2a1; SERCA2, Atp2a2) in control or TKO mice maintained at room temperature or shifted to 4°C for 1 hr (Figure 1—figure supplement 2A and B). Similarly, there was no change in myoregulin (Mrln) (Figure 1—figure supplement 2C). Although there was a small apparent increase in sarcolipin (Sln) mRNA in the control WT mice shifted to 4°C, there was no statistical difference with the TKO mice (Figure 1—figure supplement 2D). Together, these data are consistent with the absence of a cell-autonomous effect in either thermogenic adipocytes or skeletal muscle.

To unravel these unexpected findings, we next undertook a pharmacological approach to address the ability of the TKO mice to resist hypothermia. Treatment of control and TKO mice with the selective SERCA inhibitor cyclopiazonic acid (CPA) (Wang et al., 2021) reduced core body temperature at room temperature (Figure 2A). As expected, CPA treatment resulted in a faster rate of decline and to a lower extent when the mice were exposed to 4°C (Figure 2B). However, there was no significant difference in the response to CPA between the control and TKO mice. The CPA-induced reduction in body temperature is directly correlated with observable skeletal muscle paralysis, consistent with skeletal muscle contraction activity as an important component in thermal regulation (Blondin and Haman, 2018), and further indicates that SERCA functions downstream of TIGAR function.

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Since Tigar deficiency in neither adipocytes nor skeletal muscle recapitulated the protection of acute hypothermia but that paralysis results in a rapid reduction in core body temperature, we next assessed whether the cold resistance of the TKO mice was due to shivering thermogenesis (ST). To address this, we next treated mice with the highly specific skeletal muscle nicotinic acetylcholine receptor competitive antagonist tubocurare (Bowman, 2006). Control male mice treated with 0.4 mg/kg tubocurare became relatively inactive with essentially no detectable locomotor activity at room temperature 10 min post injection that lasted for another 20–30 min (see Figure 2—video 1). Over this time period, there was a concomitant decline in core body temperature that began to recover as the effects of tubocurare began to wear off (Figure 2C). Tubocurare treatment of the TKO male mice at room temperature also displayed a similar decline and recovery of body temperature but that remained somewhat higher than the control mice (Figure 2C). However, shifting the male mice to 4°C following tubocurare injection resulted in a rapid decline in body temperature such that by 30 min the control mice’s body temperature was reduced to less than 25°C (Figure 2D). In contrast, the tubocurare-injected TKO male mice only display a reduction in body temperature to approximately 34°C under identical conditions. Moreover, while the tubocurare-treated control mice placed at 4°C remained completely immobile, the TKO mice display apparent normal locomotor activity (Figure 2—video 1). Female mice are more sensitive to tubocurare than male mice (Maurya et al., 2013). The body temperature in WT female mice kept declining in 1 hr period after a lower dose (0.15 mg/kg) tubocurare injection at ambient temperature. Nevertheless, the decline of body temperature in the TKO female mice was resistant to the effects of tubocurare (Figure 2E).

The tubocurare dose-response for male mice is shown in Figure 2F, with the control WT mice having an LD₅₀ of 0.44 mg/kg whereas the TKO mice having an LD₅₀ of 0.55 mg/kg (Figure 2F). Injection of a lethal tubocurare dose (0.60 mg/kg) initially resulted in 50% if the control mice display a loss of voluntary locomotor activity by 150 s, but this required 240 s in the TKO mice (Figure 2G). At 10 min, 100% of the control mice were dead, whereas it took nearly 25 min for 100% of the TKO mice to die (Figure 2H). The tubocurare resistance of the TKO mice was not due to the changes in expression of the nicotinic acetylcholine receptor subunits with no significant differences in the skeletal muscle expression of the α1 (Chrna1), β1 (Chrnb1), or γ (Chrng) subunits (Figure 2—figure supplement 1A–C) and with a small reduction in the d (Chrnd) and ε (Chrne) subunits (Figure 2—figure supplement 1D and E) in the TKO mice. These data suggest that the protection against hypothermia was due to cholinergic signaling to the skeletal muscle.

Consistent with increased skeletal muscle energetic demand, ATP levels were significantly decreased with an apparent increase in ADP and AMP in the TKO mice after 1 hr at 4°C (Figure 3A). Stable isotope metabolic flux assessment with H₂O¹⁸ demonstrated increased incorporation of orthophosphate into ATP, ADP, and AMP in the TKO mice at 4°C, which was not significantly different at room temperature (Figure 3B). Plasma creatine was also increased twofold in the TKO at 4°C (Figure 3C), reflecting the increased phosphocreatine turnover (CP M4 phosphate fragment m/z 83, Figure 3D). The appearance of creatine in the plasma is likely due, in part, to the increased rate of shivering that is known to induce skeletal muscle leakage (Baird et al., 2012).

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Raw ¹⁸O enrichment data of stable isotope metabolic flux assessment with H₂¹⁸O was collected in quadricep white muscle of both ambient temperature housed and 4°C 1 hr exposed wild type (WT) and whole-body Tigar knockout (TKO) mice, as described in ‘Method details’.

The data was used in Figure 3.

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The mechanics of energetic supply at 4°C was further examined by using a widely targeted LC/MS/MS protocol (Hoshino et al., 2012). Key pathways found to be significant were pentose phosphate pathway, purine nucleotide cycle, and amino acid utilization pathways (Figure 3—figure supplement 1A). The pentose phosphate pathway cycle fills the purine nucleotide cycle that acts in concert with the malate-aspartate shuttle and the TCA cycle to preserve cellular energetics (Hatazawa et al., 2015). In particular, glycine/serine, histidine, and methionine metabolism also drives the input of the tricarboxylic acid cycle intermediates serving as anapleurotic inputs (Figure 3—figure supplement 1B). Together, these interactions serve to recycle AMP generated by increased skeletal muscle energy demand.

Mitochondria are a primary source of both ATP production and heat generation during muscle contraction. To determine whether mitochondrial oxidative phosphorylation (OXPHOS) efficiency and/or capacity are altered in TKO mice, permeabilized skeletal muscle fiber bundles were prepared from red portions of the gastrocnemius muscle from the control and TKO mice immediately after 1 hr cold exposure. ADP-stimulated respiratory kinetics (i.e., K_m and V_max) were virtually identical between WT and TKO muscle during respiration supported by carbohydrate (Figure 3—figure supplement 2A and B) and/or lipid-based substrates. Rates of ATP production (JATP) and oxygen consumption (JO₂) measured simultaneously under clamped submaximal and maximal ADP-demand states were also similar in muscle from the control and TKO mice (Figure 3—figure supplement 2C and D), yielding similar ATP/O efficiency ratios (Figure 3—figure supplement 2E). Finally, in vitro studies of the EDL muscle showed no differences in force-frequency, peak specific tension, or time to one-half relaxation between the control and TKO mice (Figure 3—figure supplement 2F and G). Together, these data indicate that the skeletal muscle contraction-induced thermogenesis in TKO mice is not due to any intrinsic change in skeletal muscle mitochondrial efficiency of contractile function and therefore must result from a change in signaling that in turn drives an enhanced skeletal muscle response.

Since the TKO mice are resistant to tubocurare and the major driver of skeletal muscle contraction is cholinergic signaling, we next generated cholinergic neuron-specific Tigar knockout (chTKO) mice using the choline acetyl-CoA transferase ChAT-Cre mice (Rossi et al., 2011). As the superior cervical ganglion (SCG) is composed of approximately 80% cholinergic neurons (Wang et al., 1990; Morales et al., 1995; Juranek and Wojtkiewicz, 2015), we collected the fresh SCGs (Figure 4—figure supplement 1) and confirmed that Chat^Cre resulted in the loss of Tigar in cholinergic neurons by immunofluorescence co-localization of TIGAR with the vesicular choline transporter (VChAT) (Figure 4A) and by immunoblotting analyses of TIGAR protein in the SCGs (Figure 4B). Similar to the TKO mice, the chTKO mice were also resistant to acute cold exposure compared to the control Chat^Cre mice (Figure 4C). Treatment with the SERCA inhibitor CPA, at room temperature (Figure 4D) or at 4°C (Figure 4E), also resulted in paralysis and a decline in core body temperature that was not significantly different between the Chat^Cre and chTKO mice, although the reduction in core body temperature was much greater at 4°C than at room temperature. The chTKO mice were also resistant to tubocurare at room temperature with both less of a core body temperature drop and a faster recovery (Figure 4F). In agreement with the TKO mice (Figure 2D), the resistance to cold-induced hypothermia by acute 4°C challenge was very dramatic in the chTKO mice compared to the control mice (Figure 4G). Similarly, the chTKO mice were highly resistant to the paralytic actions of tubocurare (Figure 4—video 1). Moreover, immunoblotting of the SCG for the neuron activation marker c-Fos further demonstrated that the control mice displayed a small increase in c-Fos protein levels when cold challenged that was further increased in the chTKO mice (Figure 4H). As controls, there were no significant differences in the protein levels of the vesicular acetylcholine transporter (VAChT) or the plasma membrane choline transporter (ChT) proteins. These data demonstrate that the cholinergic neuron TKO mice recapitulate the cold resistance and pharmacological characteristics of the whole-body TKO mice, consistent with these mice displaying increased cholinergic tone.

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The superior cervical ganglion (SCG) tissues were collected and snap-frozen in liquid nitrogen from the Chat^Cre and cholinergic neuron-specific Tigar knockout (chTKO) mice, as shown in Figure 4—figure supplement 1 for SCG dissection.

The SCGs from the three mice of the same strain were pooled and processed by beads homogenization to acquire tissue lysates. 15 μg of the tissue lysate were used for TIGAR (30 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section. The left four lanes of the raw image (lysate from the mice at ambient temperature) were used for Figure 4B to confirm the efficiency of TIGAR protein loss in SCG of the chTKO mice. A detailed description of the raw images is shown in Source data 1.

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Figure 4—source data 2

The superior cervical ganglion (SCG) tissues were collected and snap-frozen in liquid nitrogen from the Chat^Cre and cholinergic neuron-specific Tigar knockout (chTKO) mice, as shown in Figure 4—figure supplement 1 for SCG dissection.

The SCGs from the three mice of the same strain were pooled and processed by beads homogenization to acquire tissue lysates. 15 μg of the tissue lysate were used for choline acetyltransferase (ChAT, 70 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section. The left four lanes of the raw image (lysate from the mice at ambient temperature) were used for Figure 4B. A detailed description of the raw images is shown in Source data 1.

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The superior cervical ganglion (SCG) tissues were collected and snap-frozen in liquid nitrogen from the Chat^Cre and cholinergic neuron-specific Tigar knockout (chTKO) mice, as shown in Figure 4—figure supplement 1 for SCG dissection.

The SCGs from the three mice of the same strain were pooled and processed by beads homogenization to acquire tissue lysates. 15 μg of the tissue lysate were used for actin β (42 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section. The left four lanes of the raw image (lysate from the mice at ambient temperature) were used for Figure 4B. A detailed description of the raw images is shown in Source data 1.

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Figure 4—source data 4

The superior cervical ganglion (SCG) tissues were collected and snap-frozen in liquid nitrogen from both ambient temperature housed and 4°C 30 min exposed Chat^Cre and cholinergic neuron-specific Tigar knockout (chTKO) mice, as shown in Figure 4—figure supplement 1 for SCG dissection.

The SCGs from the three mice of the same strain were pooled and processed by beads homogenization to acquire tissue lysates. 15 μg of the tissue lysate were used for TIGAR (30 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section. The right eight lanes of the raw image (lysate from SCG tissues) was used for Figure 4H to confirm the efficiency of TIGAR protein loss in SCG of the chTKO mice. The left two lanes of the raw immunoblotting image represent the lysates of soluble fraction (RIPA lysis buffer extracted) of whole-brain tissues from the Chat^Cre and chTKO mice, respectively. A detailed description of the raw images is shown in Source data 1.

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The superior cervical ganglion (SCG) tissues were collected and snap-frozen in liquid nitrogen from both ambient temperature housed and 4°C 30 min exposed Chat^Cre and cholinergic neuron-specific Tigar knockout (chTKO) mice, as shown in Figure 4—figure supplement 1 for SCG dissection.

The SCGs from the three mice of the same strain were pooled and processed by beads homogenization to acquire tissue lysates. 15 μg of the tissue lysate were used for c-Fos (62 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section. The right eight lanes of the raw image (lysate from SCG tissues) were used for Figure 4H to show the increase in c-Fos protein in SCG of the chTKO mice under cold exposed. The left two lanes of the raw immunoblotting image represent the lysates of soluble fraction (RIPA lysis buffer extracted) of whole-brain tissues from the Chat^Cre and chTKO mice, respectively. A detailed description of the raw images is shown in Source data 1.

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Figure 4—source data 6

The superior cervical ganglion (SCG) tissues were collected and snap-frozen in liquid nitrogen from both ambient temperature housed and 4°C 30 min exposed Chat^Cre and cholinergic neuron-specific Tigar knockout (chTKO) mice, as shown in Figure 4—figure supplement 1 for SCG dissection.

The SCGs from the three mice of the same strain were pooled and processed by beads homogenization to acquire tissue lysates. 15 μg of the tissue lysate were used for vesicular acetylcholine transporter (VAChT, 55 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section. The right eight lanes of the raw image (lysate from SCG tissues) were used for Figure 4H to show VAChT protein in the SCGs. The left two lanes of the raw immunoblotting image represent the lysates of soluble fraction (RIPA lysis buffer extracted) of whole-brain tissues from the Chat^Cre and chTKO mice, respectively. A detailed description of the raw images is shown in Source data 1.

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The superior cervical ganglion (SCG) tissues were collected and snap-frozen in liquid nitrogen from both ambient temperature housed and 4°C 30 min exposed Chat^Cre and cholinergic neuron-specific Tigar knockout (chTKO) mice, as shown in Figure 4—figure supplement 1 for SCG dissection.

The SCGs from the three mice of the same strain were pooled and processed by beads homogenization to acquire tissue lysates. 15 μg of the tissue lysate were used for choline transporter (ChT, 55 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section. The right eight lanes of the raw image (lysate from SCG tissues) were used for Figure 4H to show ChT protein in that 63 and 70 kDa bands were observed in the SCGs. The left two lanes of the raw immunoblotting image represent the lysates of soluble fraction (RIPA lysis buffer extracted) of whole-brain tissues from the Chat^Cre and chTKO mice, respectively. A detailed description of the raw images is shown in Source data 1.

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The superior cervical ganglion (SCG) tissues were collected and snap-frozen in liquid nitrogen from both ambient temperature housed and 4°C 30 min exposed Chat^Cre and cholinergic neuron-specific Tigar knockout (chTKO) mice, as shown in Figure 4—figure supplement 1 for SCG dissection.

The SCGs from the three mice of the same strain were pooled and processed by beads homogenization to acquire tissue lysates. 15 μg of the tissue lysate were used for actin β (42 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section. The right eight lanes of the raw image (lysate from SCG tissues) were used for Figure 4H to show actin β protein in the SCGs. The left two lanes of the raw immunoblotting image represent the lysates of soluble fraction (RIPA lysis buffer extracted) of whole-brain tissues from the Chat^Cre and chTKO mice, respectively. A detailed description of the raw images is shown in Source data 1.

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If increased cholinergic input is responsible for the increased energy demand and heat production by skeletal muscle, then the skeletal muscle metabolic characteristics of the TKO mice should be recapitulated in the chTKO mice. As shown in Figure 5, widely targeted LC/MS/MS in the gastrocnemius muscle of the chTKO mice identified the same three key pathways – pentose phosphate pathway, purine nucleotide cycle, and amino acid utilization pathways – that display a greater induction at 4°C. Together, these data are consistent with increased cholinergic signaling to skeletal muscle that is responsible for the increased skeletal muscle thermogenesis following cold exposure.

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Raw metabolites data were collected in the gastrocnemius muscle of both ambient temperature housed and 4°C 1 hr exposed Chat^Cre and cholinergic neuron-specific Tigar knockout (chTKO) mice as described in ‘Method details’.

The data was used in Figure 5.

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To address the basis of the apparent increase in cholinergic signaling, we first asked whether there was a change in synaptic density in the NMJ. The acetylcholine receptor aggregates in the synaptic cleft and can be specifically labeled with α-bungarotoxin (Pratt et al., 2018), as shown for the whole EDL muscle (Figure 6A). Quantification revealed a large degree of heterogeneity in the number of clustered acetylcholine receptors per muscle (Figure 6B). Nevertheless, there was a clear statistical increase in the TKO mice compared to the control mice. Consistent with the increase in synaptic junctions, the total amount of acetylcholine was also increased in skeletal muscle extracts from the TKO and chTKO mice compared to their respective control mice at both room temperature and after cold exposure (Figure 6C and D).

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The culture SH-SY5Y cells were collected from both scrambled and whole-body Tigar knockout (TKO) cells, and 30 μg of the cell lysates were used for TIGAR (30 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section.

The left four lanes of the raw image (lysate from neuroblastoma cells) were used for Figure 6E to confirm the efficiency of TIGAR protein loss in the SH-SY5Y neuroblastoma TKO cells. The right four lanes of the raw image represent the TIGAR immunoblotting of the cell lysates from 7-day differentiated neuroblastoma cells. A detailed description of the raw images is shown in Source data 1.

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Figure 6—source data 2

The culture SH-SY5Y cells were collected from both scrambled and whole-body Tigar knockout (TKO) cells, and 30 μg of the cell lysates were used for actin β (42 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section.

The left four lanes of the raw image (lysate from neuroblastoma cells) were used for Figure 6E to show actin β protein in the SH-SY5Y neuroblastoma cells. The right four lanes of the raw image represent the actin β immunoblotting of the cell lysates from 7-day differentiated neuroblastoma cells. A detailed description of the raw images is shown in Source data 1.

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As it is not possible to determine the rate of acetylcholine synthesis in vivo, we took advantage of SH-SY5Y cells that have characteristics of cholinergic neurons (Kovalevich and Langford, 2013). Using sgRNA and Cas9, we effectively knocked out Tigar in several clonal SH-SY5Y cells, two representative control, and sgRNA knockout cells as shown in Figure 6E. The amount of acetylcholine secreted into the medium over a 24 hr period was increased in the TKO SH-SY5Y cells compared to controls (Figure 6F). Labeling with [1,2]-¹³C glucose demonstrates an increase in the m2-labeled acetylcholine levels in the TKO SH-SY5Y cells (Figure 6G). Similarly, labeling with U-¹³C glucose also demonstrated an increase in m2-labeled acetylcholine in the TKO SH-SY5Y cells (Figure 6H). Moreover, there was a threefold increase in the enrichment of the C2-C4 fragment, m1-glutamate m/z 129, indicating that the acetyl group in acetylcholine originated from glycolysis via pyruvate dehydrogenase (PDH) (Figure 6I, Figure 6—figure supplement 1A and B). This is because the m2 acetyl group formed via PDH generates ¹³C atoms at positions 4 and 5 of glutamate, whereas the m2 ¹³C glutamate (m/z 130) at positions 2 and 3 is derived from pyruvate carboxylase (PC)-generated oxaloacetate (Madhu et al., 2020) schematically shown in Figure 6—figure supplement 1A. Analyses of the m/z 128–130 C2-C4 glutamate fragment distinguish these isotopomers with the enrichment of m1 glutamate (129 fragment) approximately five times more abundant than the m2 glutamate (130 fragment) in the TKO cells (Figure 6—figure supplement 1B).

If the changes in glucose flux occurred through increased flux through PDH, then we should also observe a concomitant increase in the levels of acetyl-CoA that is directly generated by PDH from pyruvate. As shown in Figure 6—figure supplement 1C, acetyl-CoA levels were increased approximately 1.5-fold and acetyl-carnitine levels approximately 2.5-fold (Figure 6—figure supplement 1D). Since acetyl-CoA is converted to acetyl-carnitine in the mitochondria, these demonstrate that Tigar deficiency not only increases glycolysis but also results in increased mitochondria uptake of acetyl-CoA (via PDH). The increase in mitochondria PDH flux would in turn drive an increase in oxygen consumption. Seahorse flux analyses of oxygen consumption rate (OCR) clearly demonstrated that the TKO SH-SY5Y cells had higher rates of OCR compared to the control SH-SY5Y cells (Figure 6—figure supplement 1E). Together, these data demonstrate that Tigar deficiency in SH-SY5Y neuroblastoma cells results from an increase in intracellular acetyl-CoA levels consistent with increased glycolysis and direct conversion of pyruvate to acetyl-CoA by PDH, as opposed to PC. It is the increase in PDH flux into the tricarboxylic acid cycle that drives increased mitochondria oxidation.

A generalized increased cholinergic tone would be expected to reduce blood pressure and heart rate but with increased skeletal muscle shivering. To assess these physiological parameters, we used implanted telemetry probes to simultaneously determine body temperature, mean arterial pressure (MAP), heart rate (beats per minute [BPM]), and skeletal muscle electromyography (EMG) in the cold-exposed Chat^Cre and chTKO mice (Figure 7). Consistent with the rectal temperature measurements previously observed, the telemetry probe also demonstrated that the chTKO mice are more resistant to cold-induced hypothermia than Chat^Cre mice (Figure 7A). At room temperature, there was no significant difference in blood pressure, but following cold exposure the blood pressure of the control mice increased, whereas the blood pressure of the chTKO mice decreased such that by 60 min the chTKO mice had a blood pressure of 25 mmHg less than the control mice (Figure 7B). Similarly, the heart rate was not statistically different at room temperature but increased substantially more in the control mice than in the chTKO mice following cold exposure (Figure 7C). As these data strongly suggest increased cholinergic signaling to enhance skeletal muscle contraction, we utilized EMG to directly determine skeletal muscle activity. A representative image of EMG tracers and root mean square (RMS) derivation is shown for the Chat^Cre and chTKO mice when shifted to 4°C for 30 min (Figure 7D). Quantification of these types of data as the integral of RMS during the time course of cold exposure demonstrated that at room temperature and following 10 min of cold exposure there is no significant difference in skeletal muscle contraction activity between the Chat^Cre and chTKO mice (Figure 7E). However, the contraction activity of the control Chat^Cre mice plateau at this level, whereas the chTKO mice shivering activity further increases to nearly twice the level of the control mice.

Figure 7

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Figure 7—source data 1

The extensor digitorum longus (EDL) muscles were collected from both ambient temperature housed and 4°C 1 hr exposed Chat^Cre and cholinergic neuron-specific Tigar knockout (chTKO) mice, and 30 μg of the tissue lysate were used for AMPKα (62 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section.

The raw image was used for Figure 7F. A detailed description of the raw images is shown in Source data 1.

https://cdn.elifesciences.org/articles/73360/elife-73360-fig7-data1-v2.zip

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Figure 7—source data 2

The extensor digitorum longus (EDL) muscles were collected from both ambient temperature housed and 4°C 1 hr exposed Chat^Cre and cholinergic neuron-specific Tigar knockout (chTKO) mice, and 30 μg of the tissue lysate were used for phospho-^Thr172AMPKα (62 kDa) immunoblotting analysis as described in the ‘Immunoblotting’ section.

The raw image was used for Figure 7F. A detailed description of the raw images is shown in Source data 1.

https://cdn.elifesciences.org/articles/73360/elife-73360-fig7-data2-v2.zip

Download elife-73360-fig7-data2-v2.zip

The increased skeletal muscle shivering associated with increased ATP turnover and energy expenditure would be expected to result in a compensatory increase in AMP-dependent protein kinase (AMPK) activity. As shown in Figure 7F, there was no significant difference in the activation site phosphorylation (T172) of the AMPK α subunit in the EDL muscle at room temperature between the control Chat^Cre and chTKO mice. However, after 1 hr at 4°C, the control mice display the expected increase in T172-AMPKα phosphorylation that was further increased in the chTKO mice. Together, the relative changes in cold--induced blood pressure, heart rate, skeletal muscle shivering, and increase in AMPK activation are fully consistent with the chTKO mice displaying a generalized increase in cholinergic signaling following cold exposure.

Currently, there is a substantial effort to understand and develop approaches to increase energy expenditure through the development/activation of brown/beige adipose tissue thermogenesis. A fraction of adult humans can respond to cold stress by inducing the expression of thermogenic adipose tissue under relatively long-term cold acclimation in adult humans (McNeill et al., 2021). However, it remains somewhat controversial if this inducible thermogenic human brown adipose tissue is the rodent equivalent of beige or brown adipocytes (Cannon et al., 2020; Samuelson and Vidal-Puig, 2020; Virtanen and Nuutila, 2021). In addition, whether the mass of the human-inducible thermogenic adipocytes is sufficient to significantly contribute to cold-responsive thermogenesis for core temperature maintenance or weight loss has also been questioned (Blondin et al., 2014; Virtanen and Nuutila, 2021). In contrast, skeletal muscle accounts for approximately 40% of healthy body mass and is the dominant factor for basal metabolic rate and the major driver of energy expenditure during physical exercise (Haman et al., 2010; Blondin et al., 2014). In rodents, non-shivering thermogenesis (NST) is driven by UCP1-dependent mitochondrial uncoupling in both brown and beige adipocytes by the creatine phosphorylation/dephosphorylation cycle in beige adipocytes (Cohen and Spiegelman, 2015; Kazak et al., 2015; Jung et al., 2019). There are also reports of a skeletal muscle-based NST via a sarcolipin-dependent futile cycle of SERCA-dependent calcium transport in the sarcoplasmic reticulum (Bal et al., 2012; Maurya et al., 2015). In humans, maximal NST can increase energy production equivalent to about two times above the resting metabolic rate (RMR), while moderate ST is 2.5–3.5 times RMR and maximal shivering has been measured at about five times RMR (Haman et al., 2010; Blondin et al., 2014). In contrast, exercise at ~50% VO₂ max generates approximately 15 times RMR (Weber and Haman, 2005; Blondin et al., 2014). While a substantial effort has focused on NST, shivering and physical activity are the dominant forms of involuntary and voluntary heat production during acute cold exposure. Thus, it is essential to develop a deeper understanding of the physiological integrated role of skeletal muscle with thermogenic adipose tissue to have a complete and detailed molecular understanding of energy balance and its implications for potential interventions to promote and maintain weight loss.

The data presented in this article demonstrate that cholinergic neuron deficiency of Tigar results in an enhancement of cold-induced skeletal muscle contraction/shivering that is responsible for an increase in heat generation and protection of acute cold-induced hypothermia. This response is likely mediated by an increase acetylcholine signaling at the NMJ, consistent with the parallel induction of tubocurare resistance and increased cholinergic neuron activation. Interestingly, despite the increase in synaptic density and acetylcholine levels of mice at room temperature that are mildly cold stress (thermoneutrality for mice is approximately 29–31°C; Ganeshan and Chawla, 2017), Tigar deficiency does not appear to significantly affect body temperature, skeletal muscle contraction activity, or the overall skeletal muscle metabolite profile at room temperature. However, following acute cold challenge there are marked changes in these parameters between WT and cholinergic TKO mice including a metabolic profile indicative of increased ATP turnover. These findings suggest that there is an increase in cholinergic signaling reserve that only becomes apparent when cholinergic signaling demand is high.

In addition to the well-established role of sympathetic activation to increase brown and beige adipocytes thermogenesis (Zhu et al., 2019), both cholinergic and sympathetic activation can affect vascular blood flow and behavorial activities that also contribute to body temperature regulation (Nakamura et al., 2022). Although we cannot rule out the contribution of vasoconstriction to the conservation of core body temperature under cold exposure, heat loss from the mouse tail only accounts for 5–8% of whole-body heat loss (Skop et al., 2020) and is therefore unlikely to play a significant role in the thermoregulation observed by cholinergic Tigar deficiency.

In addition to TIGAR as a small carbohydrate phosphatase, previous studies have shown that TIGAR can directly interact with hexokinase 2 and the HOIP subunit of the linear ubiquitin assembly complex (Cheung et al., 2012; Tang et al., 2018). Thus, although it is formally possible that TIGAR increases cholinergic signaling due to alterations in protein-protein interactions, we believe that it is more likely a result of increased glycolysis due to its phosphatase function that is occurring in the cholinergic neurons as this would result in an increase in acetyl-CoA levels. The steady-state acetyl-CoA concentrations in neuronal mitochondrial and cytosol are several folds lower than the acetyl-CoA Km values for choline acetyltransferase. More specifically, following cholinergic differentiation there is a further decrease in acetyl-CoA levels that results in a further substrate dependence of acetylcholine biosynthesis (Ronowska et al., 2018). In contrast, several studies have also shown that choline uptake via the choline transporter is rate limiting for acetylcholine synthesis (Lockman and Allen, 2002). Although we have not measured choline uptake activity, Tigar deficiency had no significant effect on the amount of choline transporter protein. However, metabolic analyses clearly demonstrated an increased glucose-derived acetyl moiety into acetylcholine and increased steady-state acetylcholine levels with increased acetyl-CoA and acetyl-carnitine levels.

Thus, the simplest and most direct mechanism for the increase in acetylcholine levels induced by Tigar deficiency is the increased formation of acetyl-CoA through enhanced glycolysis, which in turn leads to increased production of acetylcholine in cholinergic neurons. At present, it is not possible to distinguish the amount of synaptic acetyl-CoA from skeletal muscle acetyl-CoA at the NMJ, but future studies using spatial mass spectroscopy (MALDI) may be able to resolve this issue.

Finally, it is also important to recognize that myasthenia gravis is a neuromuscular disease caused by autoantibodies against components of the NMJ and in particular the nicotinic acetylcholine receptor (Gilhus et al., 2019). Current therapeutic strategies are focused on the use of steroids or immune suppressants to block the immune system and drugs to increase the transmission of acetylcholine signaling such as acetylcholine esterase inhibitors (Gilhus et al., 2019). Similarly, central cholinergic neurons undergo severe neurodegeneration in Alzheimer’s disease and agents that enhance cholinergic signaling, particularly cholinesterase inhibitor therapies, providing significant symptomatic improvement in patients with Alzheimer’s disease (Ferreira-Vieira et al., 2016; Ahmed et al., 2019). Interestingly, recent reports show that the NMJs are stable in human age-related sarcopenia and in patients with cancer cachexia, which are different from the rodent models in that the sarcopenia is linked with the endplate fragmentation in NMJ (Jones et al., 2017; Boehm et al., 2020). As such, inhibition of TIGAR function may also provide a novel approach to improve skeletal muscle function, memory, and dementia through increased cholinergic signaling while limiting the deleterious metabolic consequences secondary to currently available therapies.

All data generated or analysed during this study are included in the manuscript and supporting file; Source Data files have been provided for Figures 1, 3, 4, 5, 7, Figure 1-figure supplement 1, 3, and 6.

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Author details

1. Yan Tang

   Departments of Medicine, Albert Einstein College of Medicine, Bronx, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Software, Supervision, Validation, Visualization, Writing – original draft, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-1377-422X

2. Haihong Zong

   Departments of Medicine, Albert Einstein College of Medicine, Bronx, United States

   Contribution

   Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Resources, Software, Supervision, Validation, Visualization, Writing – original draft

   Competing interests

   No competing interests declared

3. Hyokjoon Kwon

   Departments of Medicine, Albert Einstein College of Medicine, Bronx, United States

   Contribution

   Data curation, Formal analysis, Investigation, Methodology, Resources, Software, Validation, Visualization, Writing – original draft

   Competing interests

   No competing interests declared

4. Yunping Qiu

   Departments of Medicine, Albert Einstein College of Medicine, Bronx, United States

   Contribution

   Data curation, Formal analysis, Investigation, Methodology, Resources, Software, Supervision, Validation, Visualization, Writing – original draft

   Competing interests

   No competing interests declared

5. Jacob B Pessin

   Departments of Medicine, Albert Einstein College of Medicine, Bronx, United States

   Contribution

   Formal analysis, Investigation, Methodology, Software, Validation, Writing – original draft

   Competing interests

   No competing interests declared

6. Licheng Wu

   Departments of Medicine, Albert Einstein College of Medicine, Bronx, United States

   Contribution

   Data curation, Formal analysis, Investigation, Methodology, Resources, Software, Validation, Visualization

   Competing interests

   No competing interests declared

7. Katherine A Buddo

   East Carolina Diabetes and Obesity Institute and the Department of Physiology, Brody School of Medicine East Carolina University, Greenville, United States

   Contribution

   Data curation, Formal analysis, Investigation, Methodology, Resources, Software, Validation

   Competing interests

   No competing interests declared

8. Ilya Boykov

   East Carolina Diabetes and Obesity Institute and the Department of Physiology, Brody School of Medicine East Carolina University, Greenville, United States

   Contribution

   Data curation, Formal analysis, Investigation, Methodology, Resources, Software, Validation

   Competing interests

   No competing interests declared

9. Cameron A Schmidt

   East Carolina Diabetes and Obesity Institute and the Department of Physiology, Brody School of Medicine East Carolina University, Greenville, United States

   Contribution

   Data curation, Formal analysis, Investigation, Methodology, Resources, Software, Validation

   Competing interests

   No competing interests declared

10. Chien-Te Lin

    East Carolina Diabetes and Obesity Institute and the Department of Physiology, Brody School of Medicine East Carolina University, Greenville, United States

    Contribution

    Data curation, Formal analysis, Investigation, Methodology, Resources, Software, Validation

    Competing interests

    No competing interests declared

11. P Darrell Neufer

    East Carolina Diabetes and Obesity Institute and the Department of Physiology, Brody School of Medicine East Carolina University, Greenville, United States

    Contribution

    Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Resources, Software, Supervision, Validation, Visualization, Writing – original draft

    Competing interests

    No competing interests declared

12. Gary J Schwartz

    1. Departments of Medicine, Albert Einstein College of Medicine, Bronx, United States
    2. Departments of Neuroscience, Albert Einstein College of Medicine, Bronx, United States
    3. The Fleischer Institute of Diabetes and Metabolism, Albert Einstein College of Medicine, Bronx, United States

    Contribution

    Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Resources, Software, Supervision, Validation, Writing – original draft

    Competing interests

    No competing interests declared

13. Irwin J Kurland

    1. Departments of Medicine, Albert Einstein College of Medicine, Bronx, United States
    2. The Fleischer Institute of Diabetes and Metabolism, Albert Einstein College of Medicine, Bronx, United States

    Contribution

    Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Resources, Software, Supervision, Validation, Writing – original draft

    Competing interests

    No competing interests declared

14. Jeffrey E Pessin

    1. Departments of Medicine, Albert Einstein College of Medicine, Bronx, United States
    2. The Fleischer Institute of Diabetes and Metabolism, Albert Einstein College of Medicine, Bronx, United States
    3. Departments of Molecular Pharmacology, Albert Einstein College of Medicine, Bronx, United States

    Contribution

    Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Supervision, Validation, Visualization, Writing – original draft, Writing – review and editing

    For correspondence

    jeffrey.pessin@einsteinmed.edu

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0003-2041-2726

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