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Introduction

Cre-loxP technology has been widely adopted as a tool for selective cell-targeted manipulation of gene expression [1]. Through the use of the Cre-loxP system in mice, cell/tissue-specific promoters drive Cre recombinase expression in the organ of interest, following which Cre recombinase can excise the floxed gene, allowing for generation of a tissue-specific gene knockout (KO) mouse [1]. In addition, many Cre-loxP systems now employ inducible systems, such as those that are sensitive to tamoxifen but insensitive to endogenous oestrogen, thereby allowing the generation of tissue-specific, temporally controlled gene KO adult mice. Of importance, we have become increasingly aware that Cre recombinase expression or activity per se can impact tissue function. This can arise from the design of the construct used to drive Cre expression, as certain Cre constructs incorporate additional genes, such as the human growth hormone mini-gene as a polyadenylation sequence and recently reported in beta cells using the mouse insulin promoter to drive Cre expression [2]. Conversely, Cre expression driven by the myosin heavy chain-α promoter to generate cardiac myocyte-specific gene KOs produces a transient heart failure phenotype [3]. These studies highlight important considerations to be taken into account when using Cre-loxP systems to study gene/protein function, in particular the use of the appropriate control littermate for comparison. Here, we report whether similar considerations apply to the tamoxifen-inducible human α-skeletal actin (HSA) promoter to drive Cre expression in skeletal muscle.

Methods

Animal care

All animals received care according to the Canadian Council on Animal Care and all procedures were approved by the University of Alberta Health Sciences Animal Welfare Committee. All animal husbandry was performed by University of Alberta Health Sciences Laboratory Animal Services staff. Six-week-old HSA-Cre male mice [Jackson Laboratory, Bar Harbor, ME, USA; stock no. 025750; Tg(ACTA1-cre/Esr1*)2Kesr] and their wild-type (WT) littermates (Jackson Laboratory, stock no. 000664, C57BL/6J) received daily i.p. injections with either vehicle (corn oil) or tamoxifen (50 mg/kg) for 5 days. All mice were allowed a 1 week washout post-tamoxifen, following which 8-week-old mice were placed on either a low-fat diet (LFD) (10% kJ from lard; Research Diets, New Brunswick, NJ, USA, D12450J) or a high-fat diet (HFD) (60% kJ from lard; Research Diets D12492) for 12 weeks. Animals were euthanised at 20 weeks of age via i.p. injection of sodium pentobarbital (12 mg) following a 2 h fast. No randomisation was carried out and experimenters were not blinded to group assignment and outcome assessment.

Cell culture

C2C12 skeletal muscle myoblasts (ATCC, Rockville, MD, USA) were cultured and differentiated into myotubes as previously described [4]. Following 4 days of differentiation, C2C12 myotubes underwent Lipofectamine-mediated transfection with either a control or Cre recombinase-expressing plasmid (Addgene, Cambridge, MA, USA, plasmid no. 13775) for 24 h. Media was replaced with serum-free media for 24 h and collected. INS-1 832/3 cells (referred to, herein, as INS-1) were cultured and used for the assessment of glucose-stimulated insulin secretion (GSIS) as previously described [5]. As a positive control for stimulating GSIS, separate groups of INS-1 cells were treated with D-Ala2 glucose-dependent insulinotropic polypeptide (D-Ala2 GIP).

Assessment of glucose homeostasis

I.p. glucose (2 g/kg body weight) and insulin (0.7 U/kg body weight) tolerance tests were performed in 16- and 17-week-old overnight-fasted WT and HSA-Cre mice, respectively. Blood glucose measurements were performed via tail bleed with the Accu-Chek Advantage system (Roche, Indianapolis, IN, USA) in blood samples obtained at the end of the fast (0 min), followed by blood samples at 15, 30, 60, 90 and 120 min following glucose/insulin administration. Plasma was extracted from whole blood collected from mouse tails for measuring insulin levels as previously described [6].

Whole-body in vivo metabolic assessment

Indirect calorimetry was performed in 18-week-old WT or HSA-Cre mice using Oxymax metabolic cages (Columbus Instruments, Columbus, OH, USA). Animals were initially acclimatised in the system for 24 h, following which the subsequent 24 h was used for data collection (e.g. oxygen consumption) as previously described [6].

Determination of muscle triacylglycerol (TAG) levels

Frozen powdered gastrocnemius muscle tissue was extracted in a 2:1 chloroform:methanol solution, following which the supernatant phase was retained for the assessment of TAG content with an enzymatic assay kit (Wako Pure Chemical Industries, Osaka, Japan) as previously described [6].

Western blotting

Protein samples were prepared from powdered frozen gastrocnemius tissue (20 mg) homogenised in protein lysis buffer, following which they were subjected to western blotting protocols as previously described [7]. Akt and phospho (p)-Akt (9272S and 4060L); glycogen synthase kinase (GSK)3β and p-GSK3β (5676S and 9331L); mammalian target of rapamycin (mTOR) and p-mTOR (2983S and 5536); and Cre recombinase (15036) antibodies were purchased from Cell Signaling (Danvers, MA, USA) and prepared in a 1:500 dilution in 5% bovine serum albumin (BSA).

Real-time PCR analysis

First-strand cDNA was synthesised from total RNA using the SuperScript III synthesis system (Invitrogen, Carlsbad, CA, USA). Real-time PCR was subsequently carried out using TaqMan gene expression assays (Applied Biosystems, Foster City, CA, USA) for quantification of relative mRNA transcript levels (see electronic supplementary material [ESM] Table 1 for primer list) as previously described [7].

Immunohistochemistry

Pancreases were fixed in 4% formalin and embedded in paraffin, sliced (5 μm) and mounted on slides. Citrate buffer was used for antigen retrieval, and immunohistochemistry was performed using either guinea pig anti-insulin antibody (Abcam, Cambridge, MA, USA) at a 1:250 dilution or rabbit anti-glucagon antibody (Millipore, Billerica, MA, USA) at a 1:5000 dilution in 5% BSA. Antibodies were visualised using the 3,3′-diaminobenzidine colorimetric system and images were captured using a Leica DMRE microscope (Leica, Wetzlar, Germany) as previously described [8].

Nuclear/cytoplasmic fractionation

Nuclear and cytoplasmic protein fractions from gastrocnemius tissue were extracted using the Nuclear Extract Kit (Active Motif, Carlsbad, CA) according to the manufacturer’s protocol as previously described [9]. For western blotting analyses, voltage-dependent anion-selective channel (VDAC)1 and histone H3 were used as cytoplasmic and nuclear markers, respectively.

Co-immunoprecipitation

Equal amounts of protein lysates were incubated with anti-Cre antibody and a non-specific IgG antibody as a negative control overnight at 4°C. To pull down the antigen–antibody complex, 40 μl protein G Dynabeads (Thermo Fisher Scientific, Waltham, MA, USA) were added to all samples and incubated at 4°C for 2 h. The beads were washed three times with ice-cold PBS, and bound proteins were eluted by boiling at 95°C for 5 min with 20 μl of SDS protein loading buffer, following which they were subsequently processed for western blotting as previously described [9].

Statistical analysis

No samples were excluded from our data analyses. All values are presented as means ± SEM. Significant differences were determined using an unpaired, two-tailed Student’s t test or one- or two-way ANOVA followed by Bonferroni post hoc analysis.

Results

Glucose tolerance is improved in HSA-Cre mice

We observed no differences in body weight or composition between WT and HSA-Cre mice fed either a LFD or HFD for 12 weeks (Fig. 1a, b). However, while HSA-Cre mice fed a LFD demonstrated no change in glucose tolerance and circulating insulin levels, robust improvements in both variables were observed in HFD-fed HSA-Cre mice (Fig. 1c, d).

Fig. 1
figure 1

Improved glucose tolerance in HSA-Cre mice. (a) Body weight (n = 6), (b) body composition (n = 3), (c) glucose tolerance (n = 13) and (d) circulating insulin levels (n = 5–6) in tamoxifen-treated WT and HSA-Cre mice supplemented with either a LFD or HFD for 8 weeks. (e, i) Insulin tolerance (e, n = 5; i, n = 9) and (f–h, j–l) insulin signalling (Akt, GSK3β and mTOR phosphorylation) in gastrocnemius muscle (n = 4–5) from tamoxifen-treated WT and HSA-Cre mice supplemented with a LFD (e–h) or HFD (i–l) for 9 weeks. (m) Insulin and glucagon staining of pancreatic islets from tamoxifen-treated WT and HSA-Cre mice supplemented with a LFD (n = 4). Scale bars, 10 μm. (n) mRNA profiling of key genes regulating pancreatic function in pancreases from tamoxifen-treated WT and HSA-Cre mice supplemented with a LFD (n = 3–4). FC, fold change. (o) Glucose tolerance (n = 6) and (p) circulating insulin levels (n = 3–5) in corn-oil-treated WT and HSA-Cre mice supplemented with either a LFD or HFD for 8 weeks. (q) Insulin tolerance in corn-oil-treated WT and HSA-Cre mice supplemented with either a LFD or HFD for 9 weeks (n = 6). White bars/squares, LFD WT mice; light-grey bars/triangles, LFD HSA-Cre mice; dark-grey bars/squares, HFD WT mice; hatched bars/black squares, HFD HSA-Cre mice. Values are means ± SEM. Differences were determined using two-way ANOVA, followed by Bonferroni post hoc analysis. *p < 0.05 vs WT counterpart

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Insulin tolerance and signalling are not changed in HSA-Cre mice

WT and HSA-Cre mice fed a LFD exhibited similar insulin sensitivity during an i.p. insulin tolerance test (Fig. 1e), whereas phosphorylation of canonical members of the insulin signalling pathway (Akt, GSK3β, mTOR) was similar in gastrocnemius muscles following a 2 h fast (Fig. 1f–h). These indices of insulin sensitivity were also similar between WT and HSA-Cre mice fed a HFD (Fig. 1i–l). Importantly, insulin and glucagon staining-based immunohistochemistry revealed no discernible differences in islet morphology, whereas mRNA expression of key genes regulating pancreatic function was also similar between tamoxifen-treated WT and HSA-Cre mice (Fig. 1m, n). In addition, we could not detect mRNA expression for Cre recombinase in pancreatic (islet) tissues of HSA-Cre mice (data not shown).

Glucose tolerance in the absence of tamoxifen-mediated liberation of Cre recombinase

To demonstrate that the improved glucose tolerance in HSA-Cre mice is dependent on tamoxifen-mediated liberation of Cre recombinase within skeletal muscle, we repeated our studies in mice treated with vehicle (corn oil). Corn-oil-treated HSA-Cre mice fed a LFD or HFD demonstrated no differences in glucose tolerance, displayed similar circulating insulin levels in response to glucose and had similar insulin sensitivity compared with their WT littermates (Fig. 1o–q).

Skeletal muscle-specific Cre recombinase expression

Western blotting experiments demonstrated that Cre recombinase is expressed in both corn-oil- and tamoxifen-treated HSA-Cre skeletal muscles (gastrocnemius and quadriceps), but not the liver or heart (Fig. 2a). Because Cre recombinase in the HSA-Cre mouse is bound to a complex with a mutated oestrogen receptor and heat shock protein (Hsp)90, we next demonstrated, using co-immunoprecipitation, that Cre recombinase was still associated with Hsp90 in corn-oil-treated HSA-Cre mice, whereas no association between Cre recombinase and Hsp90 was observed following treatment with tamoxifen (Fig. 2b). Likewise, tamoxifen treatment of HSA-Cre mice resulted in the nuclear translocation of Cre recombinase (Fig. 2c).

Fig. 2
figure 2

Cre recombinase expression and nuclear translocation in HSA-Cre mice. (a) Cre recombinase expression in gastrocnemius muscle, quadriceps muscle, heart and liver from WT and HSA-Cre mice treated with corn oil (Corn) or tamoxifen (TMX) (n = 4). (b) Co-immunoprecipitation (IP) demonstrating that Cre recombinase remained bound to Hsp90 in HSA-Cre mice treated with corn oil, whereas TMX treatment liberated Cre recombinase from its mutated oestrogen receptor complex with Hsp90 (n = 3). (c) Nuclear and cytosolic fractions were prepared from gastrocnemius muscles from TMX-treated HSA-Cre mice, and the presence of Cre recombinase was seen only within the nuclear fraction (n = 3). Cre was normalised to VDAC1 (cytosol [cyto]) or histone 3 (nuclear [Nuc]). Values are means ± SEM. *p < 0.05, unpaired, two-tailed Student’s t test

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Substrate preference and energy expenditure in HSA-Cre mice

Indirect calorimetry studies in tamoxifen-treated, HFD-fed mice at 18 weeks of age showed similar respiratory exchange ratios, whole-body oxygen consumption rates, ambulatory activity, and water and food intake between genotypes (Fig. 3a–e). Following euthanasia of mice, we assessed TAG content in gastrocnemius muscle and mRNA expression profiles for key regulators of energy metabolism/mitochondrial function, since improved mitochondrial function often elicits improvements in glucose homeostasis [10]. However, we observed no differences in gastrocnemius muscle TAG content between WT and HSA-Cre mice (Fig. 3f), nor did we observe changes in mRNA expression for a number of genes involved in mitochondrial function (Fig. 3g), fatty acid metabolism/oxidation (Fig. 3h, i) or ketone body metabolism/oxidation (Fig. 3j). Conversely, non-significant increases in the genes encoding peroxisome proliferator-activated receptor (PPAR)-δ and PPAR γ-coactivator 1-α expression were observed in HSA-Cre mice (Fig. 3i; p = 0.069 and 0.068, respectively).

Fig. 3
figure 3

Characterisation of potential contributors to improved glucose tolerance in HSA-Cre mice. (a) Whole-body oxygen consumption rate (n = 5), (b) respiratory exchange ratio (RER) (n = 5), (c) ambulatory activity (n = 3), (d) food intake (n = 3) and (e) water intake (n = 4) in tamoxifen-treated WT and HSA-Cre mice supplemented with a HFD for 10 weeks. (f) Gastrocnemius muscle TAG content in tamoxifen-treated WT and HSA-Cre mice supplemented with a HFD for 12 weeks (n = 4–5). mRNA expression profiles for genes involved in (g) mitochondrial function, (h) lipolysis, (i) fatty acid oxidation and (j) ketone body metabolism in tamoxifen-treated WT and HSA-Cre mice supplemented with a HFD for 12 weeks (n = 4). Dark-grey bars/squares, HFD WT mice; hatched bars/black squares, HFD HSA-Cre mice. (k) Cre recombinase mRNA expression in C2C12 myotubes transfected with a control plasmid (light-grey bars) or a Cre recombinase-expressing plasmid construct (dark-grey bars) (n = 4). (l) Insulin secretion in INS-1 cells treated with CM from C2C12 myotubes overexpressing Cre recombinase (n = 7–8). Values are means ± SEM. *p < 0.05 vs the high-glucose only group, one-way ANOVA followed by Bonferroni post hoc analysis. Cq, quantification cycle; FC, fold change

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Conditioned media (CM) from C2C12 myotubes overexpressing Cre recombinase does not augment insulin secretion

We posited that expression/activation of Cre recombinase in skeletal muscle leads to the production of a secreted factor that improves beta cell insulin secretion in HSA-Cre mice. Accordingly, we transfected Cre recombinase into C2C12 myotubes (Fig. 3k), following which CM from these cells was applied to INS-1 beta cells during a GSIS test. CM from C2C12 myotubes overexpressing Cre recombinase failed to augment GSIS compared with CM from C2C12 myotubes transfected with a control plasmid, whereas D-Ala2 GIP robustly enhanced GSIS (Fig. 3l).

Discussion

Our findings support the notion that tissue-specific Cre recombinase expression may influence cellular/tissue function, as we demonstrate for the first time that expression/activation of Cre recombinase in skeletal muscle improves glucose tolerance. It is possible that Cre recombinase expression may not be truly confined to skeletal muscle in HSA-Cre mice, potentially explaining our unexpected findings, yet Cre recombinase was undetectable in the heart, liver and pancreas of HSA-Cre mice, ruling against leaky expression as a mechanism. The observed improvements were not related to changes in insulin sensitivity/signalling, but rather to increases in circulating insulin. Moreover, insulin levels only showed differences in tamoxifen-treated HFD-fed HSA-Cre mice during an i.p. glucose tolerance test, as fasted (Fig. 1d) and ad libitum (data not shown) insulin levels were identical between tamoxifen-treated HFD-fed HSA-Cre mice and their WT littermates. It remains unclear how Cre recombinase expression in skeletal muscle impacts insulin secretion, as our CM experiments in C2C12 myotubes overexpressing Cre recombinase failed to augment GSIS from INS-1 cells. Nonetheless, it is possible that Cre recombinase in skeletal muscle influences secreted factors to a different extent than that in C2C12 myotubes transfected with a Cre recombinase plasmid. Hence, further work is necessary to determine the mechanism(s) by which skeletal muscle Cre recombinase expression influences circulating insulin levels and/or insulin secretion. Our study was only conducted in male mice, and it remains to be seen whether female HSA-Cre mice exhibit the same phenotype.

Our observations have relevance as Cre-loxP systems are often used to identify how specific genes/proteins in skeletal muscle impact glycaemia and insulin sensitivity in experimental models of obesity. However, the vast majority of studies using Cre-loxP systems to investigate glucose homeostasis and insulin resistance have used the floxed mouse as the control littermate. Reasons for this likely include the rationale that a mouse expressing a floxed gene may produce a phenotype itself, whereas breeding strategies designed to produce both tissue-specific Cre recombinase control littermates and the corresponding tissue-specific gene KO are much more complex and met with a reduced frequency of required genotypes. Examples where ramifications may have been missed through the use of inappropriate control littermates to study muscle-specific actions include a report that used muscle creatine kinase (MCK)-Cre mice to generate mice deficient in muscle-specific carnitine palmitoyltransferase-1b [11], and another that used the non-inducible HSA-Cre mouse to generate mice deficient in muscle-specific growth factor receptor-bound protein 10 [12]. Although these studies demonstrated improvements in glycaemia during obesity [11] and ageing [12], the respective floxed mouse was used as the control littermate. Hence, the authors’ observations could potentially be due to the expression of Cre within skeletal muscle. It is also unknown whether MCK-Cre or non-inducible HSA-Cre mice exhibit a similar phenotype to the tamoxifen-inducible HSA-Cre mouse, and it will be important for future studies to determine if our results translate to these models.

Taken together, it appears that the HSA-Cre mouse is a necessary control littermate for any study using a Cre-loxP system to investigate skeletal muscle gene/protein function in the experimental setting of obesity/insulin resistance. Accordingly, any study using the floxed mouse as the sole control littermate will be unable to conclude whether an improvement in glucose tolerance in the corresponding skeletal muscle-specific gene KO is due to deletion of the gene within muscle itself, or the result of Cre recombinase activity.