The RNA-binding protein QKI controls alternative splicing in vascular cells, producing an effective model for therapy

Cardiovascular disease (CVD) is the leading cause of morbidity and mortality in the western world, and is characterised by progressive damage to and closure of blood vessels in key tissues (Roth et al., 2015). CVD is initiated by dysfunction of the vascular cells; the endothelial cells (ECs) and vascular smooth muscle cells (VSMCs) (Park and Park, 2015). The endothelial interface between the vessel and circulating blood is supported by VSMCs, playing essential roles in maintaining vascular tone and maturation of the blood vessel (Carmeliet, 2000). VSMCs are not terminally differentiated as adult cells. In disease scenarios, cell plasticity can be evoked, with phenotypic changes to VSMCs potentiating the development of CVD leading to complications such as atherosclerotic lesion rupture and postangioplasty restenosis (Owens et al., 2004; Davies et al., 2010; Hao et al., 2002; Moiseeva, 2001; Schatteman et al., 1996; Belaguli et al., 1999; Bennett et al., 2016).

Thus far, treatment for vascular diseases has been largely preventative rather than replacing diseased tissue due to limited availability of autologous tissue for transplantation. Cell reprogramming is a powerful technique that has led to the generation of induced pluripotent stem cells (iPSCs) from adult somatic cells, which can be directed towards any cell type required for therapy (Takahashi and Yamanaka, 2006).

ECs and VSMCs have been successfully generated from iPSCs (iPS-ECs and iPS-SMCs, respectively) but many of the mechanisms of differentiation are still unknown, leading to limited efficiency of derived differentiated cells (Margariti et al., 2012; Clayton et al., 2015). Studying the underlying mechanisms of vascular cell differentiation will allow the manipulation of vascular cells in the diseased state and provide novel targets for vascular therapy while teaching us the underpinning molecular mechanisms of development.

We have previously shown that embryonic stem cell differentiation into VSMCs requires signalling via histone deacetylase 7 (HDAC7), a class II histone deacetylase that is essential for tight regulation of gene expression (Zhang et al., 2010; Margariti et al., 2009; Dressel et al., 2001). Alternative splicing of HDAC7 plays a key role in SMC differentiation from pluripotent stem cells, while platelet-derived growth factor-BB (PDGF-BB) is known to enhance VSMC differentiation and regulate the balance of spliced to unspliced HDAC7 (Margariti et al., 2009; Zhou et al., 2011). HDAC7s modulates the serum response factor (SRF)–myocardin complex, which induces SMC differentiation from pluripotent stem cells (Zhang et al., 2010; Margariti et al., 2009; Wang et al., 2004; Chen et al., 2002). Further elucidation of the exact splicing regulators of HDAC7 is required to allow full understanding of HDAC7-mediated VSMC differentiation, which will permit the generation of more phenotypically mature VSMCs.

RNA-binding proteins (RBPs) have emerged in recent years as important modulators of post-transcriptional regulation in the cell, through altering mRNA splicing, stability, localisation and efficiency of translation (Brinegar and Cooper, 2016). This leads to changes in countless cellular process, with recent studies uncovering the roles of RBPs in the maintenance of pluripotency and commitment to cell differentiation. For example, RNA-binding motif protein 3 (RBM3) was found to be vital in osteoblast differentiation (Kim et al., 2018), while heterogeneous nuclear ribonucleoprotein K (HNRNPK) has been found to regulate myoblast proliferation and differentiation (Xu et al., 2018). Concurrently, dysregulation of RBP function is implicated in a number of diseases such as diabetes and vascular dysfunction (Yang et al., 2018).

The RBP Quaking (QKI), a member of the signal transduction and activation of RNA (STAR) family of proteins, and its various isoforms have been demonstrated to be key in vascular development (Li et al., 2003; Noveroske et al., 2002). QKI transcription begins from one major start site but alternative splicing leads to the production of three protein isoforms: QKI-5, QKI-6 and QKI-7. Each isoform has a unique C-terminal sequence, while sharing a common RNA-binding sequence, or Quaking response element (QRE) within the body of the protein (Ebersole et al., 1996; Kondo et al., 1999; Galarneau and Richard, 2005).

QKI was originally attributed as having a role in post-natal myelination of the central nervous system, while subsequent experimentation uncovered a role for QKI in blood vessel development. Mouse embryos with mutant QKI showed embryonic lethality at embryonic day (E)10–12.5 with only primitive vascular networks present due to impaired EC differentiation and an inability to recruit and differentiate mural cells (Noveroske et al., 2002).

QKI alternative splicing isoform (QKI-5) has recently been identified by our laboratory to direct iPSC differentiation to an endothelial lineage through direct binding and stabilisation of STAT3, while iPS-ECs overexpressing QKI-5 enhanced repair in an in vivo model of hind limb ischaemia (Cochrane et al., 2017). Notably, QKI has been shown to be strongly induced in vascular injury, altering the expression of myocardin, a key driver of VSMC differentiation (van der Veer et al., 2013).

Since vascular cells are capable of derivation from a common developmental progenitor (Yamashita et al., 2000), and QKI-5 is instrumental in EC differentiation from iPSCs (Cochrane et al., 2017), we hypothesised that the alternative splicing isoform QKI-6 may play a role in facilitating key mechanisms of VSMC differentiation from iPSCs. Since QKI-6 is co-localised with spliceosomal proteins in the nucleus, it may hold a pivotal role in regulating alternative splicing controlling VSMC differentiation.

In this study, we provide robust evidence that QKI-6 is highly induced during VSMC differentiation from iPSCs, stimulated by PDGF-BB, while directly regulating the splicing of HDAC7 and driving iPS-SMC differentiation. Through exploring the key mechanisms of QKI-5 and -6 in mediating alternative splicing programmes during vascular cell differentiation, we have increased the efficiency of in vivo angiogenesis and created a combination cell therapy of QKI-5-derived ECs and QKI-6-derived VSMCs.

Mouse iPSCs seeded on collagen IV and cultured in differentiation medium (DM) supplemented with 25 ng/ml of PDGF-BB (hereafter known as DM+P), differentiated towards a typical VSMC morphology (Fig. 1A, top panels) and expressed characteristic VSMC markers such as α-smooth muscle actin (SMA, also known as ACTA2) and calponin (Fig. 1A, bottom panels). In mouse iPS-derived VSMCs (miPS-SMCs), SMA was significantly induced at the mRNA level from day 4 of differentiation (Fig. 1B) with increasing calponin and SM22 expression demonstrated at the protein level (Fig. 1C).

Since VSMCs and ECs have the developmental capability to derive from a common Flk-1+ progenitor cell (Yamashita et al., 2000), it was important to assess possible co-differentiation of ECs. No CD144 (also known as CDH5) or von Willebrand factor (vWF) protein was present, as demonstrated by western blotting and immunohistochemistry, highlighting there was a robust differentiation towards VSMC fate (Fig. 1C,D). Our laboratory has recently reported that the RBP QKI-5 holds a key role in EC differentiation, angiogenesis and neovascularisation (Cochrane et al., 2017). Since the field of RBPs is fast emerging, and RBPs are now recognised as powerful, versatile regulatory units, which play pivotal roles in the regulation of cell differentiation (Guallar and Wang, 2014), we hypothesised that QKI alternative splicing may play a role in determining the developmental fate of the vascular cell. While QKI-5 expression did not increase above control levels during SMC differentiation, the level of QKI-6 was significantly induced over time at the mRNA and protein levels (Fig. 1B,C). It has been reported that myoblasts must initially express QKI-5 before being able to express any further QKI alternative splicing isoform due to the role of QKI-5 in promoting the accumulation and alternative splicing of the QKI gene (Fagg et al., 2017). In our study, we demonstrate that QKI-5 and -6 exhibit similar levels of expression early in VSMC differentiation with levels of QKI-6 then exceeding QKI-5 at later time points, indicating that QKI-6 plays a role in VSMC differentiation (Fig. 1C) (Fagg et al., 2017). In day 6 differentiated miPS-SMCs, QKI-6 was expressed alongside the VSMC markers SM22 and calponin as shown by immunofluorescence staining (Fig. 1E; Fig. S1). It was also clearly demonstrated that QKI-6 expression is significantly influenced by the presence of PDGF-BB in the culture medium. miPS-SMCs were differentiated for 24 h in the absence of PDGF-BB before 10 or 25 ng/ml PDGF-BB was added to the medium for 40 h. Both 10 and 25 ng/ml PDGF-BB led to a significant induction in QKI-6 mRNA expression in comparison to a non-treated control (Fig. 1F).

Further experiments were conducted to evaluate the role of QKI-6 in the differentiation of VSMCs. QKI-6 was overexpressed by lentiviral gene transfer (Ex-QKI-6) in day 4 differentiated miPS-SMCs for 48 h, which enhanced VSMC marker expression such as calponin, myosin heavy chain (MHC; herein, MYH11) and SMA at the mRNA level in comparison to the empty vector control miPS-SMCs (Ex-mCherry) (Fig. 2A). This data was confirmed in VSMCs overexpressing QKI-6 on day 1 of differentiation for 48 h, indicating QKI-6 is capable of inducing SMC differentiation from miPSCs (Fig. S2). Moreover, the induction of QKI-6, calponin and SM22 (also known as TAGLN) were confirmed at the protein level through western blotting and immunofluorescence staining (Fig. 2B,C; western blot quantification can be found in Fig. S3A). We previously reported that the transcription factor known to play a key role in angiogenesis, STAT3, is regulated by QKI-5 during EC differentiation. Interestingly, STAT3 was significantly reduced on QKI-6 overexpression indicating that VSMC differentiation is preferentially taking place (Fig. 2A). Quantification of cells successfully expressing the mCherry control or QKI-6 overexpression vectors alongside the smooth muscle cell marker SM22 indicated an increase in VSMC differentiation efficiency from 73% to 92% (Fig. 2C). Importantly, QKI-6 overexpression on day 3 of differentiation for 48 h also led to transcriptional activation of SM22 as shown by luciferase assay (Fig. 2D), suggesting QKI-6 enhances miPS-SMC differentiation via transcriptional activation of VSMC markers such as SM22.

In contrast, when QKI expression was knocked down through shRNA transfection (shQKI) on day 3 of miPS-SMC differentiation for 72 h, expression of the VSMC markers SMA and SM22 were significantly reduced at the mRNA and protein level in comparison to what was found in cells transfected with a non-targeting control shRNA (shNT) (Fig. 2E,F). These findings indicate that QKI-6 has a role in VSMC differentiation from iPSCs and that it could be used to enhance the efficacy of this process.

We have previously reported that PDGF-BB regulates the transcription and subsequent splicing of HDAC7, which has a role in directing the differentiation of embryonic stem cells (ESCs) to VSMCs via the SRF–myocardin complex (Margariti et al., 2009). Splicing of HDAC7 leads to the exclusion of a 57-base-pair intron, removing three stop codons and allowing transcription to begin from an alternative splice site. This process leads to a 22 amino acid longer HDAC7 spliced isoform (HDAC7s). The expression of the shorter, unspliced HDAC7 (HDAC7u) readily leads to the degradation of myocyte enhancer factor 2c (MEF2c) via the proteasome, suppressing VSMC differentiation. By contrast, the HDAC7s variant localises to the nucleus, where it binds to SRF and recruits myocardin, which is known to be a key step in SMC differentiation (Margariti et al., 2009). In this study, we have found that PDGF-BB regulates QKI-6 expression and promotes VSMC differentiation. The next question was whether QKI-6, as an RBP, could regulate HDAC7 signalling and splicing.

QKI-6 overexpression on day 1 of differentiation for 48 h led to a significant induction of HDAC7, SRF and myocardin expression at the mRNA (Fig. 3A) and protein level (Fig. 3B; western blot quantification can be found in Fig. S3B), while QKI knockdown at day 3 of differentiation for 72 h significantly reduced SRF expression (Fig. 3C). Induction of HDAC7 splicing during differentiation of miPSCs to SMCs was confirmed by qRT-PCR (Fig. 3D; splicing specific primer detection detailed in Fig. S4), but studies to date do not address the exact splicing regulators of HDAC7.

It is known that QKI can have fundamental roles in splicing alone (van der Veer et al., 2013), or in combination with a range of splicing factors, such as polypyrimidine track-binding protein (PTB; also known as PTBP1) during muscle cell differentiation (Hall et al., 2013). Hence, a screen was carried out in early differentiating cells for the induction of a range of alternative splicing factors, including QKI-5 and QKI-6. A number of genes were significantly downregulated between day 2 and day 4 of miPS-SMC differentiation (Fig. 3E). Notably, Hnrnpa2b1, known to play a role in SMC differentiation from ESCs was downregulated. Hnrnpa2b1 has been shown to peak between day 0 and 3 of SMC differentiation followed by a decline in expression, which could be the decrease we are observing in this study (Wang et al., 2012). Hnrnpf is known to be repressed by QKI-6 in oligodendrocytes and a similar response appears to be occurring in differentiating miPS-SMCs (Mandler et al., 2014). Of the Srsf isoforms, Srsf1 has been investigated for its role in neointimal hyperplasia, driving excessive VSMC proliferation. As the cells redifferentiate to a contractile state, Srsf1 levels are reduced (Xie et al., 2017). As the miPS-SMCs differentiate and become more contractile, Srsf1 would be expected to decrease as observed. Interestingly, QKI-6 was the only transcript found to be significantly induced during this process (Fig. 3E), therefore we sought to identify the association between QKI-6 and HDAC7 splicing.

Overexpression of QKI-6 was shown to enhance endogenous expression of both HDAC7 isoforms as shown by conventional PCR (Fig. 4A). cDNA was also subjected to qRT-PCR and a significant induction of HDAC7 splicing was observed in QKI-6 overexpressing cells in comparison to mCherry-transfected control cells (Fig. 4B). Similarly, upon knockdown of QKI, splicing of HDAC7 was reduced in comparison to cells transfected with non-targeting control and a significant drop in HDAC7 splicing was determined by qRT-PCR (Fig. 4C,D). In these experiments, overexpression of QKI-6 was carried out on day 1 of differentiation in order to manipulate early splicing events, while knockdown was employed on day 3 of differentiation, immediately prior to maximum levels of HDAC7 expression being reached within the cells (Fig. 3D). HDAC7 splicing increases during miPS-SMC differentiation (Fig. 3D), explaining why baseline levels of spliced HDAC7 vary in Fig. 4A and C.

To verify that QKI-6 acts upstream of HDAC7 to impact on VSMC differentiation from miPS-SMCs, HDAC7 was knocked down on day 5 of differentiation followed by QKI-6 overexpression on day 6. Cells were harvested on day 8 of miPS-SMC differentiation. Upon QKI-6 overexpression, a significant increase of HDAC7, its splicing and calponin expression were observed, whereas following knockdown of HDAC7, HDAC7 and calponin expression were significantly reduced. When HDAC7 was knocked down, the effects of QKI-6 overexpression on HDAC7 splicing and calponin expression were ablated (Fig. 4E).

As all QKI isoforms share the same KH RNA-binding domain and only differ at their C-terminus, we wished to assess that the observed effect on HDAC7 splicing was in fact QKI-6 specific. QKI-5 was overexpressed in day 3 differentiating miPS-SMCs cultured in DM+P and harvested 48 h after transfection. QKI-5 was unable to induce the expression of VSMC markers calponin or SM22 while levels of HDAC7 splicing were significantly reduced. In addition, endothelial cell markers were not observed (Fig. 4F).

These findings provide strong evidence that QKI-6 specifically acts upstream of HDAC7 and QKI-6 must act through the splicing of HDAC7 to subsequently drive SMC differentiation from iPSCs. QKI-5 and QKI-6 may also compete for binding to the HDAC7 transcript and recruit different machinery to impact mRNA turnover and splicing.

Precisely how QKI-6 interacts with HDAC7 to direct its splicing during miPS-SMC differentiation remains unknown. We therefore screened the HDAC7 mRNA sequence for the conserved QKI-binding motif ‘ACUAAC’ using the RBPmap software (Galarneau and Richard, 2005; Paz et al., 2014). A conserved QKI-binding site within intron 1 of HDAC7 at the genomic location chr15:97842906 was identified (Fig. 5A), intron 1 being where HDAC7 splicing occurs. To identify whether QKI could bind directly to this site to modulate splicing, primers incorporating the binding site were designed (Fig. 5B) and RNA immunoprecipitation was carried out. miPS-SMCs transduced to overexpress QKI-6 on day 4 of differentiation for 48 h underwent pulldown with IgG control or anti-QKI-6 antibody. RNA was extracted from the resultant RBP–mRNA complex and conventional PCR was undertaken. QKI-6 pulldown showed strong amplification of the HDAC7 intron 1 binding site. Fig. 5C is a representative image of three independent experiments. Relative band intensity was quantified across all three experiments and showed significantly increased binding of QKI-6 to HDAC7 intron 1 versus IgG control (Fig. 5C,D). Interestingly, the conserved QKI-binding motif was also identified in intron 1 of the human HDAC7-212 isoform. An RNA-binding assay was carried out in 293T cells overexpressing QKI-6 for 48 h and QKI was again seen to directly bind to this location (Fig. S5).

To understand the functional role and potential applications of QKI-derived vascular cells, a series of in vitro and in vivo functional assays were carried out. Day 3 miPS-SMCs transfected with QKI-6 for 48 h exhibited enhanced contractile capabilities compared to controls, indicating more functional cells (Fig. 6A). In order to study the tube-forming capacity of the cells, ECs and SMCs differentiated from iPSCs were transfected with a QKI-5 or QKI-6 overexpression vector, respectively, on day 3 of differentiation, and vascular tube formation assessed after 48 h of overexpression. QKI-derived vascular cells had significantly increased segment tube length and branch number in comparison to controls (Fig. 6B). Persistence of QKI-derived vascular structures was not assessed in comparison to controls, but due to the added stability VSMCs provide to vessels, we would hypothesise that these structures would have increased longevity in culture and would be less likely to degrade as quickly as controls or ECs alone.

In vivo, cells underwent QKI overexpression as described above and Matrigel plug assays on the flanks of C57BL6 mice demonstrated that both control (mCherry) and QKI-derived miPS-SMCs and miPS-ECs produced a significantly greater number of vascular structures per image than the PBS (vehicle) control. Remarkably, there was a significant increase in number of vascular-like structures present when QKI-5- and QKI-6-derived miPS-ECs and miPS-SMCs were combined versus mCherry-derived miPS-vascular cells (Fig. 6C,D). Immunofluorescence staining demonstrated incorporation of injected cells into vessel-like structures, successfully expressing CD144 and SM22 in vivo (Fig. 6E; Fig. S6).

VSMCs are highly heterogeneous cells and are derived from numerous developmental locations (Majesky, 2007). The origin of VSMCs has been shown to impact greatly on the development of vascular diseases such as atherosclerosis (Bennett et al., 2016; DeBakey and Glaeser, 2000). Here, VSMCs can act as protective stabilisers of the atherosclerotic plaque, while others dedifferentiate, proliferate and induce apoptosis in response to the local environment (Libby et al., 2011; Gomez and Owens, 2012). This can lead to plaque instability, rupture and further complications after therapeutic intervention such as in-stent restenosis (Qian et al., 2007; Alfonso et al., 2014). Other conditions that have a microvascular component, such as diabetes, are known to be potentiated by VSMC regression and loss (Montero et al., 2013; Beltramo and Porta, 2013).

By understanding the developmental programming of VSMCs through investigation of transcriptional and epigenetic mechanisms of differentiation from iPSCs, we can identify novel therapeutic targets and produce innovative cell therapies for ischaemic disease.

Alternative splicing of mRNA allows the production of several protein products from one gene locus (Yabas et al., 2015). This largely impacts on the scale of genetic diversity and can occur in a cell- or tissue-specific manner in response to environmental cues. Alternative splicing allows for fine-tuning of complex cellular responses such as cell differentiation in health and disease (Nilsen and Graveley, 2010; Black, 2003; Modrek et al., 2001). In 2002, Noveroske et al., and in 2003 Li et al. detailed lethal blood vessel defects and loss of vessel integrity in mouse embryos null or deficient for the RBP QKI, respectively. These effects were due to loss of the QKI-5 splice site, resulting in an inability to remodel the yolk sac vasculature, a combination of poor EC development and a deficiency of VSMC recruitment and maturation (Li et al., 2003; Noveroske et al., 2002). This demonstrates the importance of QKI alternative splicing in successful embryonic development. In recent years, the mechanistic details of how QKI regulates splicing during differentiation has begun to be investigated. Fagg et al. detailed cross-regulation of the QKI isoforms in directing differentiation of myoblasts, while van der Veer et al. have shown that QKI is capable of regulating myocardin expression after arterial damage of adult SMCs (van der Veer et al., 2013; Fagg et al., 2017). Owing to the role of QKI in adult and embryonic vascular cells, we hypothesised that QKI could be involved in the differential fate of vascular cells.

iPSC technology has revolutionised the future of regenerative medicine therapy and disease modelling, but to maximise the impact of this for CVD therapy, a thorough understanding is required of the post-transcriptional mechanisms of iPSC differentiation. Here, we show how alternative splicing of the RBP QKI during differentiation of iPSCs to vascular cells results in a cell fate decision directed by the choice of key growth factors supplied to the cells, which direct cell differentiation towards a specific vascular cell lineage such as VSMCs.

We recently reported how during EC differentiation from iPSCs, under the control of vascular endothelial growth factor (VEGF), QKI-5 acted to stabilise VE-cadherin expression and VEGFR2 transcriptional activation, increasing the efficiency of differentiation and leading to increased neovascularisation and angiogenesis (Cochrane et al., 2017). On addition of PDGF-BB during differentiation of miPS-SMCs, QKI-6 is selectively induced and directly impacts on alternative splicing of HDAC7, resulting in a greater SMC differentiation potential. We show that in early differentiation of SMCs, QKI-5 is expressed but this is maintained at low levels while QKI-6 continues to be induced. This demonstrates the multi-layered complexity of iPSC differentiation and how alternative splicing can be harnessed to alter cell fate decisions.

RBPs are part of highly complex systems and require tight regulation of their cellular localisation, expression levels, translation efficiencies and splicing events for normal cell development and stability (Baralle and Giudice, 2017). Particularly during cell differentiation, tightly orchestrated alternative splicing events occur, which are highly sensitive to the cellular environment and other stimuli. Previously and in this study, we have demonstrated how the presence of VEGF induces EC fate and PDGF-BB directs VSMC fate to differentiating iPSCs. These culture conditions also implicate QKI splicing events inducing the specific splicing isoform required for precise vascular cell development. This demonstrates one of the multiple regulatory layers of cell differentiation and lineage commitment (Wang et al., 2008; Fu and Ares, 2014).

QKI-6 is capable of driving HDAC7 splicing and SMC gene expression while its absence eradicated this ability. Adding further PDGF-BB during differentiation was found to be integral in inducing the expression of QKI-6, linking the growth factor to the previous finding that PDGF-BB regulates levels of spliced HDAC7 within VSMCs. QKI-6 regulated HDAC7 splicing by directly binding to the splicing region of HDAC7 in intron 1. Furthermore, QKI-6 was unable to direct splicing or SMC differentiation to the same extent in the absence of HDAC7 expression, emphasising the important role of the mechanism of QKI-6-mediated HDAC7 splicing in VSMC differentiation.

Neff et al. have previously discussed how VSMCs are often forgotten in the generation of tissue engineered blood vessels. Although ECs are the main functional unit of the blood vessel, SMCs contribute to a much larger portion of the vessel wall and are crucial in development and remodelling of the vasculature along with maintaining vascular structure and tone in response to physiological cues (Neff et al., 2011; Benjamin et al., 1998; Lilly, 2014). Therefore, it is logical that using ECs alongside VSMCs when studying vascular development will lead to improved understanding of vascular physiology. On combination of QKI-derived miPS-ECs and miPS-SMCs, we saw increased angiogenic potential in vitro and in vivo, in single and co-culture conditions. This demonstrates a basis for using the QKI splicing isoforms in vascular therapy.

In conclusion, this study elucidates a novel role for the RBP QKI-6 and provides further insight into the splicing mechanisms of HDAC7 during SMC differentiation from miPSCs. We have demonstrated mechanisms of vascular cell differentiation from iPSCs in which QKI alternative splicing in response to lineage-specific growth factors is crucial. QKI-5 binds to the 3′-UTR of STAT3, leading to its stabilisation and phosphorylation, resulting in VEGFR2 activation and VE-cadherin stabilisation. This is mediated upstream by VEGF-induced expression of ETS-1, promoting QKI-5 function (Cochrane et al., 2017). Conversely, PDGF-BB leads to the induction of QKI-6 and iPSC differentiation towards a VSMC fate. QKI-6 binds directly to intron 1 of HDAC7, causing a 57-base-pair intron to be excised, and a splicing event to occur. β-catenin is then capable of inducing SMC proliferation while spliced HDAC7, in conjunction with SRF and myocardin, activate and drive expression of SMC genes such as SM22 and calponin, therefore driving a VSMC fate (Zhang et al., 2010; Margariti et al., 2009; Zhou et al., 2011; Yang et al., 2016) (Fig. 7). We envisage that these iPS-derived vascular cells could be employed for tissue engineering, disease modelling and drug screening in a patient-specific manner prior to administration to the diseased patient. This will lead to more effective, efficient and personalised cell sources for cardiovascular cell therapies.

Cell culture medium, serum and cell culture supplements were purchased from the ATCC, Millipore and Thermo Fisher Scientific.

Antibodies against calponin [ab46794; 1:2000 for western blotting (WB), 1:500 for immunocytochemistry (ICC)], SM22 (ab14106; 1:1000, WB; 1:200, ICC), QKI-6 (ab9906, 1:500, ICC), QKI-5 (ab9904; 1:500, WB; 1:200, ICC), GAPDH (ab125247; 1:1000, WB) were purchased from Abcam. Antibodies against SRF (sc-335, 1:1000, WB), myocardin (H-300; 1:1000, WB) and vWF (C-20; 1:50, ICC) antibodies were obtained from Santa Cruz Biotechnology. A second QKI-6 antibody (N182/17; 1:1000, WB) was purchased from Neuromab. CD144 (STJ96234, 1:200k ICC; 1:1000, WB) and HDAC7 (STJ93484, 1:1000, WB) antibodies were purchased from St. John's Laboratory, London, UK. Cy3-conjugated α-smooth muscle actin (C6198, 1:100, ICC) was obtained from Sigma.

The secondary antibodies for immunostaining were anti-mouse-IgG and rabbit-IgG conjugated to Alexa Fluor 568, and anti-mouse-IgG and rabbit-IgG conjugated to Alexa Fluor 488 purchased from Thermo Fisher Scientific and used at a 1:200 dilution. The secondary antibodies for western blotting were purchased from Bio-Rad and used at a 1:3000 dilution. Control shNT (sc-62166) and shQKI (sc-106468) plasmids were purchased from Santa Cruz Biotechnology and shHDAC7 from Sigma (NM_019572). The plasmids for expression of QKI isoforms 5 (217EX-T4215-Lv224) and 6 (217EX-H2552-Lv224) were designed and purchased from Genecopoeia.

Mouse iPSCs (miPSCs) were generated as previously described (Kelaini et al., 2014; Di Bernardini et al., 2014). Mouse iPSCs were cultured on gelatin (PBS; Life Technologies #10010056 containing 0.02% of gelatin from bovine skin; Sigma #G1393) in DMEM (ATCC #30-2002) supplemented with 10% fetal bovine serum (FBS) (Embryomax; Millipore #ES-009-B), 100 IU/ml penicillin and 100 μg/ml streptomycin (PenStrep) (Thermo Fisher Scientific #10378-016), 10 ng/ml recombinant human leukaemia inhibitory factor (LIF) (Millipore #LIF1010) and 0.1 mM 2-mercaptoethanol (Life Technologies #31350-010). The cells were passaged every 2 days at a ratio of 1:6. Cells were not used after passage 60. Differentiation of iPSCs towards vascular cells was induced by seeding the cells on type IV mouse collagen (5 μg/ml; R&D Systems #3410-010-01) and cells were cultured in DM containing α-MEM (Life Technologies #32571036) supplemented with 10% FBS (Invitrogen #10270106), 0.05 mM 2-mercaptoethanol, PenStrep, supplemented with 25 ng/ml PDGF (Thermo Fisher Scientific #PMG0045) for SMC differentiation or 25 ng/ml VEGF (Thermo Fisher Scientific #PMG0111) for EC differentiation for the time points indicated.

For PDGF-BB stimulation experiments, miPSCs were subjected to differentiation in the absence of PDGF-BB for 24 h. The cells were then serum starved overnight and either 10 or 25 ng/ml PDGF-BB was added to the media for 40 h. Cells were routinely tested for mycoplasma contamination fortnightly.

293T (ATCC #HEK293T) cells were cultured in DMEM (Thermo Fisher Scientific #10566016), Penstrep with 10% FBS.

Total RNA was extracted using the RNeasy Mini Kit (Qiagen #74104) and reverse transcribed by using a High Capacity cDNA Reverse Transcription Kit (Thermo Fisher Scientific #4368814) with 10 µg of RNase inhibitor (Thermo Fisher Scientific #N8080119) according to manufacturer's protocol. Resultant cDNA was diluted to a concentration of 10 ng/µl.

Relative gene expression was determined by quantitative real-time PCR (qRT-PCR), with the SYBR Green Master Mix (Life Technologies #4368702) and primers as detailed in Table S1. GAPDH served as the endogenous control. The gene was considered undetectable beyond 35 cycles.

Lentivirus was generated in 293T (ATCC #HEK293T) cells passaged to 60–70% confluency. 4.5 µg of relevant plasmid was combined with the packaging genes; 0.9 µg pCMV-dR8.2 (Addgene #8455; deposited by Bob Weinburg) and 3.6 µg pCMV-VSV-G (Addgene #8454; deposited by Bob Weinburg) with 27 µl EndoFectin Max (Genecopoeia #EF013) and 300 µl Opti-MEM 1 Reduced Serum Medium (Thermo Fisher Scientific #31985070). This was incubated for 10 min at room temperature. Cells were washed once with pure Opti-MEM which was followed by addition of 5 ml 2% FBS-Opti-MEM to the flasks. The transfection solution was added dropwise to the cells and flasks incubated overnight at 37°C. Medium was changed next morning to complete 293T medium (DMEM, Penstrep, 10% FBS). After 48 h, the first round of virus was collected and passed through a 0.45 µm filter, removing any cell debris. Medium was replaced for another 48 h when harvest process was repeated.

miPSC-derived vascular cells were allowed to differentiate for the indicated amount of time before lentiviral transduction. Equal volumes of control or overexpression/knockdown virus in DM were added to the cells with an additional 10 µg/ml of Polybrene (Santa Cruz Biotechnology #134220). Medium was changed the next morning and cells were harvested 48 h later in overexpression scenarios and 72 h after infection for knockdown.

Plasmids were transformed by combination with competent E. coli (Promega #JM109), plated on agar (Sigma, #L2897) with suitable resistance antibiotic added (Sigma, Ampicillin #BP785) and grown overnight. Plasmids from colonies picked and amplified the next day were purified using the QIAprep Spin Miniprep Kit (Qiagen #27106). DNA concentration and purity was measured by the Nanodrop spectrophotometer.

miPSCs were differentiated for the time points indicated before transfection with 500 ng–2 µg of relevant plasmid. 3 µl of EndoFectin Max (Genecopoeia #EF013) was incubated per 1 µg of plasmid in 50 µl Opti-MEM per reaction for 10 min at room temperature. A reduced volume of Opti-MEM-2%FBS was added to the culture dishes to which the transfection solution was added dropwise to the cells. After 16 h, the medium was changed back to normal differentiation medium. Cells were harvested 48 h post-transfection.

Total RNA was extracted using the RNeasy Mini Kit (Qiagen #74104) following the manufacturer's protocol. 1–2 µg RNA was reverse transcribed into cDNA using the High Capacity cDNA Reverse Transcription Kit (ThermoFisher Scientific #4368814) with 10 µg of RNase inhibitor (ThermoFisher Scientific #N8080119) according to manufacturer's protocol.

Thermocycler conditions were set at 25°C for 10 min, 37°C for 120 min, 85°C for 5 min and a 4°C hold. Resulting cDNA was diluted to a concentration of 10 ng/µl with DEPC-H₂O.

Relative gene expression was determined by qRT-PCR, using 20 ng of cDNA per sample with the SYBR Green Master Mix (Life Technologies #4368702) in a 10 μl reaction following the manufacturer's protocol. For each sample 5 μl of PCR master mix, 2 μl of primer set, 1 μl DEPC-treated water and 2 μl of cDNA (20 ng) was used. Ct values were measured using LightCycler 480 sequence detector (Roche). For each sample, PCR was performed in duplicate in a 384-well reaction plate (LightCycler 480 Roche real-time PCR plates #04729749001) using primers specific to the gene sequence (Table S1). The qPCR conditions were 5 min at 95°C for initial de-nature of the DNA followed by 40 cycles of 95°C for 15 s, 60°C for 30 s for the quantitative amplification stage with a final single cycle to create a melting curve with the conditions 95°C for 15 s, 60°C for 15 s then 95°C for 15 s. GAPDH served as the endogenous control to normalise the amounts of RNA in each sample. The gene was considered undetectable beyond 35 cycles. Primers are detailed in Table S1.

Cells were harvested and washed with cold PBS, resuspended in RIPA buffer (Sigma #R0278) and lysed by ultra-sonication (Bradson Sonifier150). The protein concentration in lysates was quantified with a Quick Start Bradford protein assay (BioRad #5000201). Lysates were normalised using 2× Laemmli sample buffer (Sigma #S3401) before 20–50 µg of lysate was applied to SDS-PAGE and transferred to Amersham Hybond P 0.45 PVDF blotting membrane (GE Healthcare #10600023). The bound primary antibodies were detected using horseradish peroxidase (HRP)-conjugated secondary antibodies (Bio-Rad #1706516, #1706515) and Clarity Western ECL Substrate (Bio-Rad #1705061).

Cell fixation was carried out with 4% paraformaldehyde and cells were permeabilised with 0.1% Triton X-100 inPBS. Cultures were blocked in 5% goat or donkey serum, followed by incubation with the desired primary antibody. Subsequent washes were followed by secondary antibody incubation and counterstaining with DAPI (Life Technologies #D1306) before mounting with Vectashield (Vector Laboratories #H-1000). Cells were imaged by fluorescence (Dmi8, Leica) and confocal microscopy (SP8, Leica).

On day 3 of miPS-SMC differentiation, cells were transfected using FuGENE 6 (Promega, # E2691) with Ex-mCherry or Ex-QKI-6 vector and the SM22 promoter (pGL3-SM22). pGL3-Luc Renilla was used as an internal control. At 48 h after transfection, luciferase and Renilla luciferase activity was detected (Promega, #E1501; #S2001). Relative luciferase Units (RLU) were defined as the ratio of luciferase activity to Renilla luciferase activity.

miPS-SMCs and 293T cells (ATCC #HEK293T) overexpressing QKI-6 were harvested at 48 h post transfection and subjected to RBP immunoprecipitation using the Magna RIP kit (Millipore, #17-700) as per the manufacturer's protocol. A QKI-6 specific antibody (Abcam, #ab9906) and rabbit IgG (included in kit) were used. Purified RNA was subjected to RT-PCR using specific primers for the predicted HDAC7 intron 1 binding site.

Day 3 differentiated miPS-SMCs were transfected with Ex-mCherry or Ex-QKI-6 for 48 h. The cells were stimulated with 40 mM KCl for 15 min and live imaged using the Nikon 6D Inverted Microscope. Fluorescently labelled cells (i.e. those successfully transfected) were measured for their cell size at time zero and 15 min. The percentage change in cell size was calculated for the duration of the imaging.

For the tube formation assays, 96-well plates were coated with growth-factor reduced Matrigel (Corning #354230). After defined treatments and time points, differentiated cells were dissociated using TrypLE (Gibco #12604013) and counted, and 30,000 cells were seeded per well with 10 µM of the ROCK inhibitor Y27632 (TOCRIS #1254) in 150 µl medium. ECs and SMCs were seeded at a ratio of 2:1, respectively. Cells were incubated for between 4 and 8 h before imaging by bright field microscopy (DMi8, Leica). Images were processed with the Angiogenesis Analyser plugin for ImageJ (Gilles Carpentier Research Web Site: Computer Image Analysis).

Animals used in these studies were all bred in-house under constant climatic conditions with free access to food and water. All experiments were performed in accordance with the Guidance on the Operation of the Animals (Scientific Procedures) Act, 1986 and approved by the Queen's University Belfast Animal Welfare and Ethical Review Body. Work was performed under the project license number PPL2821 and the Personal Licence Number 1705.

On completion of cell differentiation and treatments, cells were dissociated with TrypLE and counted using a haemocytometer. 1.5×10⁵ miPS-ECs were combined with an equal number of miPS-SMCs per treatment condition. The cells were centrifuged for 5 min at 1000 g to obtain the cell pellet. This was resuspended in 1 ml of medium and the cells were stained with 5 µl Vybrant Dil Cell Labeling Solution (Thermo Fisher Scientific #V22885) per 1 ml of cell suspension, per 10⁶ cells, for 15 min at 37°C. 2 ml of medium was then added to the suspension and the cells centrifuged for 5 min at 200 g. The resulting pellet was resuspended in 50 µl of PBS and layered on top of 200 µl Matrigel matrix basement membrane (Corning #356234). Three male 8–10-week-old C57BL6 mice were anaesthetised using 1 litre min⁻¹ of oxygen with 2% isoflurane (Attane, #12164-002-25) for induction followed by 1.5% for maintenance of anaesthesia. Hair was removed from the lower back of the mouse and the 250 µl cell/Matrigel or PBS/Matrigel vehicle control suspension was injected subcutaneously. No more than 3 plugs were injected per mouse. After 7 days, the mice were killed and the plugs were dissected from the underside of the skin. Plugs were snap frozen in liquid nitrogen before transfer into Tissue-Tek Optimal Cutting Temperature Compound (O.C.T.) (VWR #25608-930) and storage at −80°C. Plugs were sectioned using the Leica CM1900 Cryostat into 12 µm slices and placed onto Superfrost Ultra Plus Adhesion Slides (Thermo Fisher Scientific #J3800AMNT). Sections were allowed to reach room temperature for 30 min before freezing or continuing with staining.

For Haematoxylin and Eosin staining, slides were hydrated for 5 min in running distilled water to remove the O.C.T. compound. Slides were placed in haematoxylin stain for 5 min, washed for 3 min, submerged in acid alcohol for 10 s then 1% ammonia for 30 s. A 3 min eosin submersion was carried out next, followed by a serial dehydration through two 75%, 95% and 100% alcohols for 1 min each. The slides were then cleared in Xylene three times for 3 min each. Slides where then mounted with DPX and when dry, imaged by bright field microscopy using a Leica DMi8 microscope.

For immunofluorescence staining of frozen sections, slides were fixed with cold acetone for 10 min before continuing from the permeabilisation step as detailed in the above fluorescence staining section. Images were obtained using the a Leica DMi8 microscope.

Data are expressed across independent biological replicates as the mean±s.e.m. and were analysed using GraphPad Prism 5 software with a two-tailed Student's t-test for two groups or pairwise comparisons or analysis of variance (ANOVA). *P<0.05; **P<0.01; ***P<0.001 was considered significant.