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Application of ARID1A to murine formalin-fixed paraffin embedded tissue using immunohistochemistry

ARID1A is a known suppressor of tumour formation and the Human Protein Atlas antibody HPA005456 has been demonstrated in previous literature to stain tumour tissue by immunohistochemistry (IHC) in formalin-fixed paraffin embedded human tissue and human cell lines. This article details the validation of this antibody for immunohistochemistry of formalin-fixed paraffin embedded murine tissue using a Leica BondMax immunostainer. Using Western blot and IHC on murine wild-type and knockout tissue we have demonstrated that this antibody to ARID1A correctly stains murine tissue by immunohistochemistry.

Introduction
ARID1A (AT-rich interactive domain 1a) is a member of the SWI/ SNF family and its loss has been implicated as a factor in multiple premalignant and malignant conditions, including Barrett's oesophagus and oesophageal carcinoma as well as endometrial and clear cell ovarian carcinomas and their precursor endometriotic lesions [1][2][3][4] . The ARID1A antibody from Human Protein Atlas is a rabbit antibody generated against a PrEST (Protein Epitope Signature Tag) fragment of the ARID1A gene and affinity purified against the same fragment 5 . It is thus designated as being "monospecific" in that the affinity purification removes any non-specific or low affinity binders to the peptide. Through the Human Protein Atlas, the antibody has been tested on a wide variety of human tissue types and human malignancies, as well as for expression in immunofluorescence on U-2 OS, A-431 and U-251 MG cell lines. This demonstrates a nuclear expression in all cell lines and in the majority of tissue types 6 . However, the Western blot data were not supportive and did not produce staining corresponding to the expected size, although the data from a protein array did confirm a peak at the expected size. The antibody has been used to stain human colorectal cancers 7 , clear cell carcinomas 8 and on a variety of clear cell cancer cell lines 9 by immunohistochemistry.
To our knowledge, whilst the sequence homology between mouse and human ARID1A is 95%, this antibody has not been qualified using knockout tissue and has not been tested or published on murine tissue and this work represents the first data to do so.

Reagent details
Details of all reagents with reference to the immunohistochemical staining procedure can be found in Table 1.

Antibody details
Anti-ARID1A is a monospecific rabbit polyclonal generated to a PrEST sequence -PGLGNVAMGPRQHYPYGGPYDRVRTEPG IGPEGNMSTGAPQPNLMPSNPDSGMYSPSRYPPQQQQQQ QQRHDSYGNQFSTQGTPSGSPFPSQQTTMYQQQQQNYK (Table 2). The homology of the PrEST sequence used as immunogen

Amendments from Version 1
Following reviewers comments, the authors have modified the manuscript to reflect their views. The no primary antibody (NPA) data has been added to Figure 3 and an additional datafile added for the whole slide image. The authors feel that the persistant stromal staining demonstrates a limitation in the Tamoxifeninduced knockout model and not a non-specific binding of the antibody. This has been explained further in the conclusions section of the article. Donkey anti-rabbit biotin (Jackson Immunoresearch, Table 2) is specific for Rabbit IgG (Heavy and Light chains) and was affinity purified to remove cross-reactions to Bovine, Chicken, Goat, Guinea Pig, Syrian Hamster, Horse, Human, Mouse, Rat and Sheep. All slides were stained with a concentration of 4.8 µg/ml.

REVISED
Anti-GAPDH was used as a loading control for Western blots and was a rabbit monoclonal (Cell Signaling, Table 2). Detection antibody for the Western blot for ARID1a was Goat anti-rabbit IR Dye 680LT (Li-Cor Biosciences, Table 2) used at a concentration of 0.1 µg/ml and detection of GAPDH was Goat anti-rabbit IR Dye 800CW (Li-Cor Biosciences, Table 2).

Tissue details
All tissues and cell pellets used during the validation were fixed in Neutral Buffered Formaldehyde as specified (Table 3) before being transferred directly to 70% ethanol for no longer than 3 days. Tissue processing was conducted on a Leica ASP300 through graded ethanols before clearing in xylene and impregnation in molten paraffin wax (Fisher). All tissue sections were cut on a Leica rotary microtome at 3 µm.
Arid 1a -/mice were created by crossing Floxed Arid1a mice with ROSA26 Cre-ERT2 mice and resultant genotyping. Loss of Arid1a expression is expected following intraperitoneal injection of Tamoxifen.
Floxed Arid1a mice were a gift from Dr. Peri Tate, Sanger Institute, Hinxton UK; ROSA26 Cre-ERT2 mice were a gift from Prof Chambon, IGBMC, France. ES-2 cells were purchased from ATCC and RMG-II were a gift from Prof Huntsman, British Columbia Cancer Agency, Vancouver, Canada.

Experiment details
Western blot. Protein was extracted from the two clear cell carcinoma cell lines using a Tris-EDTA lysis buffer and run on a nondenaturing 3-8% Tris-acetate gel (Life Technologies). Following electrophoresis, the transfer membrane was probed with 0.2 µg/ml of anti-rabbit ARID1A (HPA005456) at 4°C overnight and 0.1 µg/ml anti-GAPDH (14C10) for the same length of time.
Detection of the anti-rabbit ARID1A was with Goat anti-rabbit IRDye 680LT (Li-Cor Biosciences) and the GAPDH was with Goat anti-rabbit IRDye 800CW (Li-Cor Biosciences) both at 0.1 µg/ml.
The antibody was validated on a Leica BondMax instrument using a Leica Intense R kit to a standardised in-house protocol. All reagents were from Leica as part of the Intense R kit and were conducted at room temperature, unless otherwise specified. All staining steps included individual washes in Leica Bond Wash after each step, as part of the protocol (Table 4). A full protocol for the validated conditions can be found in the supplementary material. In this protocol, the step named "primary" refers to the anti-ARID1a primary antibody.
A slide using the same conditions and retrieval but omitting the primary antibody was used to control for any background staining due to the retrieval and detection steps.

Results
To determine the correct cell line to utilise and to confirm the equivocal Western blot data from Human Protein Atlas, the antibody was used to stain a Western blot of two cell lines; ES-2 and RMG-II, both of which are cell lines derived from clear cell carcinoma and have been previously demonstrated as ARID1a wild-type and mutated, respectively 9 . It could be demonstrated that the HPA ARID1A antibody showed positive expression in ES-2 cell lines at the expected size of 270 kDa and no staining for RMG-II. The loading control of GAPDH showed that there were no loading issues ( Figure 1). Thus, these cell lines were chosen to be grown, formalin fixed and processed into paraffin wax for immunohistochemical validation.
For immunohistochemical validation, ES-2 and RMG-II cell lines were stained using three antigen retrieval conditions; ER1 (Sodium Citrate, pH6), ER2 (Tris/EDTA, pH9) and Enzyme 1 (Proteinase K, 100 µg/ml) at a fixed antibody concentration of 1 µg/ml. The enzyme retrieval demonstrated no nuclear signal for either ES2 or RMG-II cell pellets and was discarded for future work (Figure 2; Dataset a). The ER2 condition did demonstrate significant nuclear staining in the ES2 cell pellet with minimal background staining in the RMG-II cell pellet ( Figure 2; Dataset b). However, the staining in the ER1 condition was determined to give the best signal:noise ratio with no background cytoplasmic staining and crisp nuclear staining for the cell pellet (Figure 2; Dataset c). Control slides, omitting the primary antibody, were negative except for the ER2 condition in the RMG-II cell pellet where a weak cytoplasmic background could be seen (Figure 2; Dataset d). Thus there was minimal background inherent in the staining procedure. It was therefore determined that the antibody showed specificity for formalin-fixed paraffin embedded tissues and could be run on murine tissue.
Murine uterine tissue was used as positive control tissue samples, given the literature data on cell lines and endometrial tissue. The ER1 condition at 1 µg/ml demonstrated clean nuclear staining in the uterine epithelial compartment as well as nuclear staining of stromal cells. However, the nuclear staining in the stroma was not universal and distinct negative nuclei could be seen ( Figure 3A; Dataset e). There was no cytoplasmic or extracellular stromal background staining present and the antibody titrated successfully losing the intensity of staining, as expected (Dataset e). Following this, a concentration of 0.5 µg/ml was used for future preparations which provided clear and consistent staining in repeated batches using a different antibody lot (Dataset f). A No Primary antibody control (NPA) showed no staining in the epithelial or nuclear compartment ( Figure 3B; Dataset e).
Finally, when applied to a genetically engineered, tamoxifeninduced, Arid1a knockout mouse model, the staining in the uterine epithelium could be almost completely abrogated (Arrow, Figure 4b) when compared to the same area in a wild-type animal (Arrow, Figure 4a) with a small focal area of epithelial staining still present.
There was no effect of the KO on the staining in the stromal compartment.  Detailed legends for each dataset (Datasets a-f) can be found in the text file provided. For Version 2 an additional image was added to Dataset e.

Conclusions
It is clear from the use of ES2 and RMG-II cell lines that the Atlas Antibodies ARID1A antibody is specific for ARID1A in both Western blots and formalin-fixed paraffin embedded preparations of human origin and, coupled with the literature evidence, that it is validated in human tissue.
The staining pattern when applied to murine uterus showing a clear nuclear pattern, where there is a high level of sequence homology between the two species, is again consistent with the literature on this protein. When stained on an ARID1a knockout mouse model, the staining could be almost completely abrogated in the epithelial compartment but not in the stroma. Knockout mice generated in this manner are almost never 100% complete as in some cells recombination will not be induced due to issues such as low ligand (Tamoxifen) penetration or failure of the ligand to induce recombination, thus explaining the small focal area of epithelial staining. The difference in staining in the two compartments is also likely related to the same effect, as all other controls, such as omission of primary antibody remained negative. Thus, given the overwhelming data from other sources, it is likely the stromal staining reflects continuing Arid1a expression in this specific model system.
Therefore, in conclusion when taken in combination, it is clear that the anti-human ARID1a antibody is cross-reactive with murine tissue and can be used for this purpose.

Data availability
F1000Research: Dataset 1. Whole slide images from antibody validation of HPA005456 for immunohistochemistry -Version 2, 10.5256/f1000research.5514.d41579 10 Author contributions WH wrote and conceived the idea behind the article, IG performed the Western blots and mouse experiments and requested validation of ARID1A in murine tissue, JM performed all immunohistochemical staining.

Competing interests
No competing interests were disclosed. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

4.
This article, by Howatt validates an antibody to ARID1A in murine FFPE samples, is a good example et al. of the correct validations procedures that must be met when validating an antibody for use in tissue sections. Issues are relatively minor: What was the rationale for a starting concentration of 1μg/ml for initial validations?
There is a focal region of the KO epithelium which clearly appears to be positive. The authors would need to explain this.
Can the authors please add more discussion of the nature of the staining of the stromal cells. Do they consider this to be non-specific? Is such staining of stroma present in human tissue sections?
Control slides, omitting primary antibody, were carried out during the cell line phase of the experiments. Were such slides run on the murine tissue sections and can the results be included in the data. I note that the stromal staining was still present at selected concentration of 0.5μg/ml (dataset f) I have read this submission. I believe that I have an appropriate level of expertise to confirm that it is of an acceptable scientific standard, however I have significant reservations, as outlined above.
No competing interests were disclosed. Thank you for your comments which we agree with and have taken on board. Specifically: All of the validations conducted at the Histopathology/ISH facility start with a dilution of 1:100 from the manufacturers provided stock concentration and we perform 3 retrieval methods; Sodium Citrate pH6, Tris EDTA pH9 & Proteinase K. We titrate as appropriate following establishment of the optimal retrieval method. The manufacturers original stock concentration was 100ug/ml, hence the 1ug/ml starting concentration. They have subsequently changed their stock concentration to 200ug/ml.
We believe that the knockout model recombining a floxed Arid1a with CRE-ERT2 is not 100% complete as in some cells recombination will not be induced due to issues such as low ligand (tamoxifen) penetration or failure of the ligand to induce recombination. Thus explaining the area of focal staining in the epithelium. Similarly, although we have not investigated this further, we believe that the same problem underlies the stromal staining and is likely due to penetrance of the tamoxifen after being delivered intraperitoneally. Thus, given the wealth of data and the no primary antibody controls, now included in Fig 3, we feel that the stromal staining is not non-specific. Stromal staining does indeed occur in human tissue. All of the above have now been included in the conclusion section of the article.
Kind Regards, , Cancer Research UK Cambridge Institute, UK Will Howat Dear Dr Chalmers, Many thanks for your comments which we agree with and have taken on board. Specifically: We have modified the title to read "Application of anti-ARID1a antibody..." We have BLAST searched the protein sequence and it confirms 95% homology for the sequence used for immunisations. This has been added to the methods.
We accept that the methods detailing the Western blotting are more limited than the IHC methods, as is the case of the methods detailing the production of the knockout mouse model. However, we feel that the article was designed as an example of antibody validation for immunohistochemistry and that while western blotting provides important additional data, it is not the focus of the article. We thus feel that there is sufficient information in the details behind the western blotting to repeat the experiment, without clouding the article with the full methods.
The sources of the cells and mouse tissue have now been cited.
The lack of staining of RMG-II cells was consistent with the literature (Anglesio ) and et al. this information has been added to the results section.
The figure legend has been modified.
We believe that the focal staining is a result of incomplete recombination floxed Arid1a allele and Cre/ERT2. The resulting knockout is almost never 100% complete as in some cells recombination will not be induced due to issues such as low ligand (tamoxifen) penetration or failure of the ligand to induce recombination. Similarly, we believe that the staining in the stromal compartment is specific but represents a failure of delivery of Tamoxifen or of recombination. The data from no primary antibody (NPA) control which are all completely clean, now included in Fig 3, helps to demonstrate this. We have included these points in the conclusion.
The author contributions have been modified. Kind Regards,

Will Howat
No competing interests were disclosed. Competing Interests:


extfieldsofstudy: Biology Medicine

text:
Noonan and LEOPARD syndrome Shp2 variants induce heart displacement defects in zebrafish

Germline mutations in PTPN11, encoding Shp2, cause Noonan syndrome (NS) and LEOPARD syndrome (LS), two developmental disorders that are characterized by multiple overlapping symptoms. Interestingly, Shp2 catalytic activity is enhanced by NS mutations and reduced by LS mutations. Defective cardiac development is a prominent symptom of both NS and LS, but how the Shp2 variants affect cardiac development is unclear. Here, we have expressed the most common NS and LS Shp2-variants in zebrafish embryos to investigate their role in cardiac development in vivo. Heart function was impaired in embryos expressing NS and LS variants of Shp2. The cardiac anomalies first occurred during elongation of the heart tube and consisted of reduced cardiomyocyte migration, coupled with impaired leftward heart displacement. Expression of specific laterality markers was randomized in embryos expressing NS and LS variants of Shp2. Ciliogenesis and cilia function in Kupffer's vesicle was impaired, likely accounting for the left/right asymmetry defects. Mitogen-activated protein kinase (MAPK) signaling was activated to a similar extent in embryos expressing NS and LS Shp2 variants. Interestingly, inhibition of MAPK signaling prior to gastrulation rescued cilia length and heart laterality defects. These results suggest that NS and LS Shp2 variant-mediated hyperactivation of MAPK signaling leads to impaired cilia function in Kupffer's vesicle, causing left-right asymmetry defects and defective early cardiac development.

Missense germline mutations in PTPN11 are associated with Noonan syndrome (NS) (OMIM: 163950) and LEOPARD syndrome (LS) (OMIM: 151100), two autosomal dominant disorders. NS is a relatively common disorder that affects 1 in 1000-2000 live births. Individuals with NS are characterized by congenital heart defects, including atrial and ventricular septal defects, pulmonary stenosis and hypertrophic cardiomyopathy (HCM) (Tartaglia et al., 2001;Digilio et al., 2002). In addition, these individuals display short stature and facial abnormalities. LS is a more rare disorder (1:3500 live births) that shows a substantial overlap with the various symptoms of NS with two more clinical manifestations that are specific for LS: deafness and 'café au lait' spots on the skin (Mendez and Opitz, 1985;Legius et al., 2002). The congenital heart defects among individuals with PTPN11 mutations differ between NS and LS, in that pulmonary stenosis is most common in NS, whereas HCM prevails in LS (Allanson and Roberts, 1993;Sarkozy et al., 2008).
Interestingly, the biochemical properties of NS-Shp2 and LS-Shp2 are distinct, in that NS mutations enhance catalytic activity and LS mutations strongly reduce PTP activity (Keilhack et al., 2005;Kontaridis et al., 2006). The crystal structure of Shp2 shows that, in the absence of a binding partner, the N-SH2 domain interacts with the PTP domain and blocks the catalytic site (Hof et al., 1998). NS mutations predominantly reside in the interface between the N-SH2 domain and the PTP domain, resulting in the disruption of the closed conformation and enhanced catalytic activity of NS-Shp2 (Keilhack et al., 2005;Nakamura et al., 2009). By contrast, most LS mutations reside close to the active site and result in strongly reduced, yet detectable, catalytic activity (Keilhack et al., 2005;Hanna et al., 2006;Yu et al., 2013). How two mutations with opposite effects on catalytic activity result in syndromes withsimilar clinical symptoms is a conundrum that still needs to be resolved.
NS-Shp2 causes upregulation of the MAPK pathway (Feng, 1999). Whereas several studies show that LS mutations downregulate the level of phosphorylated ERK (Kontaridis et al., 2006;Stewart et al., 2010), the role of LS-Shp2 mutation on the MAPK pathway is still controversial (Oishi et al., 2006(Oishi et al., , 2009Edouard et al., 2010). Increased RAS/MAPK signaling is implicated in the gain-of-function phenotypes that are caused by expression of the LS Drosophila ortholog of PTPN11, corkscrew (csw) (Oishi et al., 2009), suggesting a mechanism by which NS and LS variants may result in similar phenotypes. However, NS and LS signaling are distinct, because it is well established that LS mutations, but not NS mutations (De Rocca Serra-Nedelec et al., 2012), enhance PI3K/AKT signaling in hearts of LS/+ knock-in mice (Kontaridis et al., 2006) as well as in fibroblasts isolated from individuals with LS (Edouard et al., 2010). The similarities in RAS/MAPK signaling of NS and LS variants may underlie the overlapping symptoms in NS and LS, whereas the differences in PI3K/Akt signaling may cause the differences in NS-and LS-induced developmental defects.
Shp2 knockout, knockdown and gain-of-function studies in a variety of organisms have begun to reveal the roles of Shp2 in embryonic development. Shp2-null mouse embryos die preimplantation due to defective Erk activation and trophoblast stem cell death (Yang et al., 2006). Dominant-negative Shp2 mutants disrupt gastrulation in Xenopus (Tang et al., 1995). Moreover, mouse models have been generated for the two most prevalent NS and LS mutations, Ptpn11-D61G/+ and Ptpn11-Y279C/+, respectively. These mice exhibit developmental defects, including reduced length, craniofacial abnormalities and congenital heart defects, reminiscent of the clinical characteristics of the human disorders (Araki et al., 2004;Marin et al., 2011). In particular, pulmonary stenosis and HCM are evident in the NS and LS mice, respectively. Shp2 is conserved in zebrafish and we previously generated the most common human NS and LS mutations in a cDNA encoding full-length zebrafish Shp2. Micro-injection of mRNA-encoding mutant Shp2 variants into zebrafish embryos at the one-cell stage induces gastrulation cell movement defects, as well as craniofacial and cardiac defects (Jopling et al., 2007). Furthermore, Stewart et al. used zebrafish embryos to show that the function of Shp2 in neural crest cells underlies LS pathogenesis (Stewart et al., 2010).
The role of NS and LS Shp2 variants in the development of cardiac defects is still unclear. Zebrafish has become a powerful model with which to study cardiac development in recent years (Bakkers, 2011;Lien et al., 2012). We set out to study early heart development in zebrafish embryos expressing NS and LS variants of Shp2, taking advantage of the transparency of zebrafish embryos, which facilitates time-lapse analysis of the onset and nature of the cardiac defects in vivo. Our analyses revealed impaired heart function and morphogenesis, which were highly similar in NS-and LS-Shp2-expressing embryos. Furthermore, the heart defects were accompanied by randomization of left-right asymmetry, which was probably due to impaired ciliogenesis and defective cilia function in Kupffer's vesicle that we observed in early embryos. Treatment with a MEK-inhibitor, CI-1040, prior to gastrulation rescued defective leftward heart displacement and the laterality defects in NS-and LS-Shp2-expressing embryos. Taken together, our results provide insight into the mechanism by which NS and LS Shp2 variants induced laterality defects through hyperactivation of MAPK signaling, resulting in cardiac defects during early development.

Expression of NS and LS Shp2 variants caused defects in cardiac function
To investigate the role of Shp2 variants in cardiac function, Shp2-D61G and Shp2-A462T mRNAs were injected at the one-cell stage. In order to monitor expression of mutant Shp2, we fused a green fluorescent protein (GFP)-peptide 2A sequence to the N terminus of Shp2, which is cleaved off auto-proteolytically (Kim et al., 2011) (supplementary material Fig. S1). Following expression of (mutant) Shp2, the developing heart was analyzed at 55 hpf by high-speed video recording. Kymographs of the atrium and ventricle were generated, allowing quantification of heart function, as described previously (Tessadori et al., 2012). Representative examples of kymographs of embryos expressing WT-Shp2 and Shp2-D61G are depicted in Fig. 1A-F. Quantification of the kymographs of Shp2-WT-, Shp2-D61G-and Shp2-A462T-injected embryos revealed significantly longer cardiac cycles in the Shp2 variant-expressing embryos, reflecting a reduced but regular heart rate (Fig. 1G). Other functional defects, such as an atrioventricular block or uncoupling of the two chambers, were not detected in Shp2-D61G-and Shp2-A462T-expressing embryos. These results illustrate functional cardiac defects, particularly a reduced heart rate, in embryos expressing NS and LS variants of Shp2.

Impaired heart asymmetry in embryos expressing NS and LS Shp2 variants
To further characterize the cardiac phenotype in response to expressing of NS and LS variants of Shp2, we first examined the overall morphology and regionalization of the heart at 55 hpf by in situ hybridization (ISH), using a panel of cardiac markers. At this stage, the heart chambers are completely formed in wild-type embryos and the heart undergoes looping morphogenesis. Analysis of heart shape using a myl7-specific probe (formerly known as cmlc2) revealed that the hearts of Shp2-D61G-and Shp2-A462T-expressing embryos exhibited either inverted looping (14% and 11%, respectively) or nonlooped heart (39% and 25%, respectively) in contrast to non-injected The hearts of WT-Shp2-(Shp2) and Shp2-D61Gexpressing zebrafish embryos were imaged by high-speed video recording microscopy at 2 dpf. White dotted lines through the atrium (a) and the ventricle (v) are indicated. (C-F) Ventricular (C,D) and atrial (E,F) kymographs from 2 dpf embryonic hearts. Note the longer period of the Shp2-D61G-expressing heart, compared with the WT-Shp2-expressing heart (double-headed arrow and white dotted vertical lines). (G) Quantitative analysis of the heart period in the ventricle or atrium of WT-Shp2, Shp2-D61G and Shp2-A462T expressing embryos (n indicates number). Whisker plots are depicted.
control and WT-Shp2-injected embryos ( Fig. 2A-D). Myocardial chamber specification was not affected in NS-Shp2-and LS-Shp2injected embryos, as determined by expression of ventricular myosin heavy chain (vmhc) (Fig. 2E-H) and atrium myosin heavy chain (amhc; myh6 -Zebrafish Information Network) ( Fig. 2I-L). Next, the formation of the endocardial cushions, from which the atrioventriclar valves will develop, was assessed by has2 expression. In Shp2-D61Gand Shp2-A462T-expressing embryos, has2 expression was not affected, compared with non-injected control and WT-Shp2expressing embryos ( Fig. 2M-P). Expression of this panel of markers was also assessed in embryos expressing Shp2-T73I (NS) and Shp2-G465A (LS) with similar results (supplementary material Fig. S2), indicating that all NS and LS variants induced similar defects. Taken together, these data indicate that expression of NS and LS variants of Shp2 led to a failure to undergo heart looping in a large proportion of the embryos, without inducing defects in cardiac chamber specification or endocardial cushion formation. In addition, the NS and LS variants of Shp2 induced cardiac defects to a similar extent during early development.
To investigate at which stage the cardiac defects arise in embryos expressing NS and LS variants of Shp2, myl7 expression was analyzed at different time points of heart development, particularly during cardiac fusion and heart tube elongation. Zebrafish heart formation is initiated by fusion of two bilateral pools of cardiomyocytes at the 16-somite stage (16 hpf ) into a cardiac disc at the midline of the embryo at 22 somites (20 hpf ) (Yelon et al., 1999;Bakkers, 2011). No obvious defects were detected in myl7 expression or cardiac fusion between non-injected control, WT-Shp2-, Shp2-D61G-and Shp2-A462T-expressing embryos up to 22 somites ( Fig. 3A-H). At 28 hpf, the heart in WT-Shp2expressing embryos formed a tube that extended from the midline to the area under the left eye ( Fig. 3I), which resembled normal heart displacement. By contrast, in Shp2-D61G-and Shp2-A462Texpressing embryos, the displacement of the heart is randomized: 23% of Shp2-D61G-expressing embryos and 28% of Shp2-A462Texpressing embryos had a heart on the right, whereas 34% of Shp2-D61G-expressing embryos and 24% of Shp2-A462Texpressing embryos had their heart tube at the midline ( Fig. 3J-L). Taken together, these results indicate that the first signs of cardiac abnormality in embryos expressing Shp2 variants are observed during asymmetric displacement of the heart tube.

Decreased cardiomyocyte migration speed and rotation in embryos expressing NS and LS Shp2 variants
Randomization of heart displacement may be caused by compromised cardiomyocyte migration (Ramsdell, 2005). Tg(myl7:GFP) embryos express eGFP in cardiomyocytes and were used for high-resolution confocal time-lapse imaging to assess cardiomyocyte migration. As WT-Shp2-expressing embryos were indistinguishable from non-injected control embryos in in situ hybridization experiments (Figs 2 and 3), we used WT-Shp2-expressing embryos as a control for the embryos expressing NS and LS variants of Shp2 for subsequent experiments. Single cells were tracked in WT-Shp2-, Shp2-D61G-and Shp2-A462T-expressing embryos from cardiac disc stage (21 somites, 19.5 hpf) for 4 h and their velocities and directions were determined. Consistent with observations of heart morphogenesis in wild-type embryos (de Campos-Baptista et al., 2008;Smith et al., 2008), cardiac cells moved towards the left and anterior part of the embryo in WT-Shp2-expressing embryos (n=4). Cardiomyocytes in the posterior part of the cardiac disc moved faster than anterior cells ( Fig. 4A-C). The Shp2-D61G-and Shp2-A462T-expressing embryos were imaged at 19.5 hpf, like the WT-Shp2-expressing embryos, and heart displacement was assessed subsequently at 24 hpf, allowing correlation between early migration defects and heart displacement. Embryos that displayed impaired heart displacement were selected for further analysis. It is noteworthy that none of the selected embryos displayed heart displacement to the right (Shp2-D61G, n=8; Shp2-A462T, n=8), which may be due to low penetrance of rightward heart displacement. Although cardiomyocytes in the selected embryos formed a cardiac tube, they showed reduced migration on the left. Representative images of an embryo expressing Shp2-A462T are depicted in Fig. 4D-F. Quantification of the speed of cardiomyocytes confirmed slower migration of cells in embryos expressing Shp2-D61G and Shp2-A462T, compared with WT-Shp2 (Fig. 4G). Cardiomyocytes in selected embryos expressing Shp2-D61G and Shp2-A462T did not cross the midline, but instead migrated anteriorly, which is consistent with defective heart displacement to the left. Rotation of the heart tube was quantified as described before and the observed 30°clockwise rotation of the cardiac cone in WT-Shp2-expressing embryos was consistent with the previously reported clockwise rotation in control embryos (Smith et al., 2008). Clockwise rotation was significantly decreased in embryos expressing Shp2-D61G and Shp2-A462T with impaired leftward heart displacement, compared with control WT-Shp2 (Fig. 4H). Our results indicate that directional cardiomyocyte migration and heart tube rotation is reduced in embryos expressing NS and LS Shp2 variants, which is consistent with the observed defects in heart tube displacement.

NS and LS Shp2 variants disrupt left/right asymmetry
To determine whether the loss of asymmetry in the hearts of embryos expressing NS and LS Shp2 variants is associated with disruption of overall L/R asymmetry, we analyzed the expression of a number of laterality markers. southpaw (spaw) is one of the earliest markers displaying asymmetric expression in development (Long et al., 2003). spaw was predominantly expressed in the left lateral plate mesoderm (LPM) in WT-Shp2-expressing embryos (Fig. 5A). spaw expression was affected in embryos expressing Shp2-D61G and Shp2-A462T, compared with WT-Shp2, in that 23% of Shp2-D61Gand 17% of Shp2-A462T-expressing embryos displayed bilateral spaw expression and 26% of Shp2-D61G-and 28% of Shp2-A462Texpressing embryos expressed spaw exclusively in the right LPM ( Fig. 5B-D). Next, expression of lefty2 (lft2), a downstream target of spaw, was investigated at the 22-somite stage (20 hpf ). In WT-Shp2expressing embryos, lft2 was detected in the left cardiac field in all cases (Fig. 5E). By contrast, expression of lft2 in Shp2-D61G-and Shp2-A462T-expressing embryos was randomized. 30% of the Shp2-D61G-expressing embryos expressed lft2 on the right side ( Fig. 5E-G). Similarly, in Shp2-A462T-expressing embryos, lft2 expression was detected at the right side in 42% of the embryos. spaw is required for visceral organ L/R asymmetry (Long et al., 2003), suggesting that defects in L/R asymmetry may not be limited to the heart in embryos expressing Shp2 variants. To investigate this, the position of the liver was assessed by analysis of the expression of fabp. fabp expression was randomized at 48 hpf in embryos expressing Shp2-D61G and Shp2-A462T, in that the liver developed on the opposite side in 20% of the Shp2-D61Gexpressing embryos and 24% of the Shp2-A462T-expressing embryos, or on both sides in 18% of the Shp2-D61G-and 9% of the Shp2-A462T-expressing embryos (Fig. 5H-K). In control WT-Shp2-expressing embryos, the liver always developed on the left side of the embryos. Analysis of the embryonic midline using the sonic hedgehog marker (Krauss et al., 1993) indicated that expression of Shp2-D61G and Shp2-A462T altered left-right patterning without disrupting the midline, which provides a barrier between the left and the right side (supplementary material Fig. S4). These results show that L/R patterning in the LPM is impaired in embryos expressing NS and LS Shp2 variants, which may lead to randomization of heart and gut laterality.

Ciliogenesis and cilia function in Kupffer's vesicle are impaired in embryos expressing NS and LS Shp2-variants
It is well established that Kupffer's vesicle (KV), a fluid-filled ciliated organ, is involved in the establishment of L/R asymmetry in zebrafish (Neugebauer et al., 2009;Liu et al., 2011). Through the oriented rotation of the cilia inside the KV, asymmetric expression of genes such as spaw and lft2 is established. Immunohistochemistry was carried out using an acetylated tubulin antibody to detect cilia in the KV of 10-somite stage embryos (14 hpf ) ( Fig. 6A-C). The mean number of cilia was 51.4±0.5 in WT-Shp2-expressing embryos, whereas in embryos expressing Shp2-D61G and Shp2-A462T, the number of cilia was significantly reduced to 30.4±0.7 and 28.5±0.4, respectively (n=10 embryos each) (Fig. 6D). In addition, the cilia length was significantly reduced in embryos expressing NS and LS Shp2 variants (both 3.2±0.1 μm, n=241 cilia for Shp2-D61G and n=172 cilia for Shp2-A462T, respectively) compared with control (4.2±0.1 μm, n=379 cilia for WT-Shp2) (Fig. 6E). Measurements of the area of KV indicated significant differences in lumen size among wild-type (4517±355 μm 2 , n=10), Shp2-D61G (2243±190 μm 2 , n=10) and Shp2-A462T (1914±256 μm 2 , n=10) embryos (Fig. 6F). These results suggest that disruption of L/R asymmetry in embryos expressing NS and LS variants of Shp2 was a consequence of defective KV function, resulting from impaired ciliogenesis in the KV. To determine the KV functionality directly, we analyzed the fluid flow in embryos expressing NS and LS variants of Shp2. Tracking of fluorescent beads injected into the KV lumen of wildtype Shp2-expressing embryos showed counterclockwise rotation of the beads (Fig. 6G). By contrast, tracking of the beads in Shp2-D61G-and Shp2-A462T-expressing embryos showed that the beads moved about randomly (Fig. 6H-I), indicating a loss of coordinated flow. In addition, beads tracked in Shp2-D61G and Shp2-A462Texpressing embryos showed a strongly reduced average flow velocity relative to wild-type Shp2-expressing embryos (Fig. 6M). These results indicate that ciliogenesis and cilia function is impaired in the KV of embryos expressing NS and LS variants of Shp2, resulting in defective KV function.
Early treatment with the MEK inhibitor CI-1040 rescued L/R asymmetry, heart asymmetry and KV function in embryos expressing NS and LS Shp2 variants Shp2 variants enhance RAS/MAPK signaling (Feng, 1999). Previously, we and others showed that inhibition of MEK rescued developmental defects in zebrafish embryos that were caused by expression of activators of the RAS/MAPK pathway (Anastasaki et al., 2009;Runtuwene et al., 2011). We investigated whether inhibition of MEK would also rescue the cardiac defects in embryos expressing NS and LS Shp2 variants. First, the efficacy of the MEK inhibitor CI-1040 to rescue early developmental defects was assessed. Expression of Shp2-D61G or Shp2-A462T in zebrafish embryos induced cell movement defects, resulting in elongated embryos at bud stage (10 hpf ) (Fig. 7A-C). Treatment with 0.25 μM CI-1040 at 4.5 hpf for 1 h rescued development at 10.5 hpf of embryos expressing Shp2-D61G and Shp2-A462T (Fig. 7D-F). Quantification of the epiboly defects confirmed the rescue of the embryos expressing NS and LS Shp2 variants (Fig. 7G). In addition, we confirmed that elevated pERK levels in embryos expressing Shp2-D61G and Shp2-A462T were reduced by CI-1040-mediated inhibition of MEK at 10 hpf (Fig. 7H).
Next, the effect of CI-1040 treatment was assessed on asymmetry of the heart in embryos expressing Shp2-D61G and Shp2-A462T. Treatment with 0.25 μM CI-1040 at 4.5 hpf for 1 h rescued leftward displacement of the heart at 24 hpf of Shp2-D61G and Shp2-A462Texpressing embryos (Fig. 7I,J). However, treatment of these embryos for 1 h with CI-1040 at the 10-somite stage (14 hpf) did not restore normal heart displacement in embryos expressing NS and LS Shp2 variants (Fig. 7K). Treatment with 0.25 μM CI-1040 at 4.5 hpf for 1 h largely rescued randomization of the left-right asymmetry markers and of the liver (supplementary material Fig. S5). These results Embryos were injected at the one-cell stage with mRNA encoding WT-Shp2 (Shp2), Shp2-D61G or Shp2-A462T and fixed at the stage indicated. In situ hybridization was carried out using probes specific for spaw (A-D, southpaw), lft2 (E-G, lefty2) and fabp (H-K, fatty acid binding protein, marking the liver). Representative pictures are shown of Shp2-D61G-expressing embryos. lft2 expression was observed only on the left side or on the right side. Organ asymmetry as assessed using the different markers was scored for embryos injected with WT-Shp2, Shp2-D61G and Shp2-A462T. Percentages of left, middle/bilateral and right expression of the markers are depicted. The number of embryos (n) is indicated.
indicate that inhibition of MEK at early stages rescues L/R asymmetry in embryos expressing NS and LS Shp2-variants.

DISCUSSION
In this paper, we used zebrafish to investigate defects in cardiac development in response to expression of NS and LS variants of Shp2. We found that NS-and LS-Shp2-expressing embryos showed impaired leftward heart displacement. Our results are consistent with a report that Noonan-associated mutations induce a cardiac looping defect in Xenopus (Langdon et al., 2012). Here, we provide insight into the underlying mechanism. The cardiac defects that we observed in zebrafish embryos are associated with L/R laterality defects and with defects in ciliogenesis. Expression of NS and LS variants of Shp2 in zebrafish embryos enhanced MAPK activation. The laterality defects were rescued by early treatment with CI-1040, a MEK inhibitor, indicating that the defects in heart asymmetry were caused by enhanced MAPK signaling before gastrulation.
Here, we focused on the cardiac defects in embryos expressing NS and LS Shp2 variants. The heart defects only became evident during cardiac tube formation, the first asymmetric process that occurs in the zebrafish heart (Bakkers, 2011). Normally, the cardiac tube is displaced to the left, but in embryos expressing NS and LS Shp2 variants, heart displacement was randomized (Fig. 3). Cell movement analyses revealed a delay in cell movements, coupled with reduced cardiac tube rotation, resulting in morphologically abnormal hearts at 28 hpf (Fig. 4). Cell tracking studies showed that loss of Nodal or BMP signaling leads to similar defects (de Campos-Baptista et al., 2008;Smith et al., 2008). Nodal mutant embryos show reduced speed and directional movement of the cardiomyocytes, resulting in disruption of the leftward morphogenesis and cardiac cone rotation. Asymmetric spaw expression in the LPM is responsible for the correct displacement of the heart tube to the left (Long et al., 2003;de Campos-Baptista et al., 2008). In line with these findings, we found that expression of spaw was randomized in embryos expressing NS and LS Shp2 variants, concomitant with its downstream gene target lft2. As a consequence, left/right asymmetry was lost and leftward displacement of the heart in embryos expressing Shp2-D61G and Shp2-A462T was compromised. In addition, randomized fabp expression suggested that not only asymmetry of the heart, but also of the liver was impaired.
Left-right asymmetry is conserved across species and defects in left-right asymmetry are collectively called laterality disease (Mercola and Levin, 2001). The phenotype we observed in zebrafish embryos expressing NS and LS Shp2 variants was characterized by randomization of left/right asymmetry. It is noteworthy that laterality disease is frequently associated with congenital heart defects (CHDs) (Ramsdell, 2005). Impaired leftright patterning can affect cardiac morphogenesis, resulting in defective septation and/or double outlet right ventricle (DORV) (Ramsdell, 2005). Cardiac defects associated with laterality diseases are also reported in Shp2-D61G knock-in mice. A proportion of the heterozygous Shp2-D61G mice displays septal defects and DORV (Araki et al., 2004). However, laterality defects are not the only cause of septation defects or DORV. Other laterality defects were not reported in NS or LS Shp2 knock-in mice, and laterality defects have not been commonly reported in humans with NS and LS. Hence, it remains to be determined whether laterality defects have a role in human NS and LS.
In zebrafish expressing NS and LS variants of Shp2, we found that the underlying mechanism for randomization of L/R asymmetry is likely impaired cilia function in the KV. In particular, cilia length and KV area were associated with Shp2 variant-induced impaired fluid flow in KV and subsequent loss of asymmetry (Fig. 6). It is well established that L/R asymmetry is mediated by cilia that generate fluid flow within the KV, resulting in expression of spaw in the left side of the LPM (Nonaka et al., 1998;Essner et al., 2005). Laser-mediated ablation of KV randomizes expression of spaw and lft2 (Essner et al., 2005). Moreover, shorter cilia result in perturbed intravesicular fluid flow, leading to loss of asymmetry in the heart, brain and viscera (Ferrante et al., 2009). KV formation is completed by the 6-somite Fig. 7. Early treatment with the MEK-inhibitor CI-1040 rescued leftward heart displacement and cilia length in embryos expressing NS and LS Shp2 variants. Embryos were injected with mRNA encoding WT-Shp2 (Shp2), Shp2-D61G and Shp2-A462T at the one-cell stage. (A-F) Representative images of bud stage embryos are depicted. The shape of the NS-and LS-Shp2-expressing embryos is restored by treatment with 0.25 μM CI-1040 at 4.5 hpf for 1 h. (G) The ratio between the length of the major and minor axes was determined as quantitative measure of the epiboly defects in NS-and LS-Shp2-expressing embryos. Averages are depicted with error bars indicating s.e.m. Statistical significance was determined using Student's t-test (**P<0.01; ***P<0.001).
(H) Embryos were treated with MEK inhibitor CI-1040 (0.25 μM) for 1 h at 4.5 hpf or mock-treated with DMSO and lysed at 10 hpf. Immunoblots of the zebrafish lysates were stained using antibodies specific for pErk and Erk. The experiment was repeated three times and representative blots are depicted here. (I-K) Embryos were mock treated with DMSO (I) or with CI-1040 for 1 h at 4.5 hpf (J) or at 10 hpf (K). Heart displacement was determined in embryos following in situ hybridization with a myl7-specific probe as in Fig. 4. Histograms represent the percentage of embryos with heart displacement to the left, middle or right. The total number of embryos (n) is indicated. stage (12 hpf ) (Oteiza et al., 2008(Oteiza et al., , 2010 and the generation of the KV depends on the dorsal forerunner cells (DFCs) that are induced during the blastula period . Defects in the specification, clustering or organization of DFCs induce defective KV organogenesis and/or ciliogenesis (Essner et al., 2005;Oteiza et al., 2010). Therefore, it will be interesting to investigate the role of the NS and LS Shp2 variants in DFC biology.
Shp2 is a well-known downstream factor in FGFR signaling (Neel and Tonks, 1997). It has been reported that FGFR1 signaling regulates cilia length in zebrafish. Morpholino (MO)-mediated knockdown of FGFR1 in zebrafish reduces cilia length in KV and perturbs directional fluid flow, thus impairing L/R patterning of the embryo (Neugebauer et al., 2009). The observation that knockdown of FGFR1, a positive regulator of MAPK signaling, has similar effects to mutant Shp2-induced hyperactivation of MAPK appears to be contradictive. However, it is not uncommon that attenuation and activation of a signaling pathway have the same effect on developmental processes. FGFR1 knockdown, on the one hand, and Shp2 variant-mediated enhanced MAPK activation, on the other, have similar effects on cilia function in KV and both impair L/R asymmetry. Our results predict that ectopic activation of MAPK signaling, e.g. overexpression of (activated) MAPK, around gastrulation would also induce laterality defects.
Defective ciliogenesis was observed in the KV of NS and LS-Shp2-expressing embryos. It remains to be determined whether ciliogenesis in other organs is also affected in these embryos. It is noteworthy that deafness is associated with LS in humans. Deafness is typically associated with defective cilia in the inner ear and future work should focus on ciliogenesis in organs other than KV in NS and LS Shp2-variant-expressing zebrafish embryos.
The cardiac defects that were induced by expression of NS and LS Shp2 variants in zebrafish embryos were indistinguishable. MAPK activation was elevated at bud stage in response to expression of both Shp2-D61G (NS) as well as Shp2-A462T (LS) (Fig. 7H), which might explain why we observed similar developmental defects at early stages. It is well established that NS-Shp2 mutations enhance MAPK activation, but LS-Shp2 often does not activate MAPK signaling. Nevertheless, it is not unprecedented that LS mutations induce defects that are associated with activation of MAPK signaling. Expression of LS-Shp2 mutants in Drosophila results in ectopic wing veins and a rough eye phenotype, characteristics of increased Erk activation. This gain-of-function phenotype is similar to that of NS-Shp2 transgenic flies (Oishi et al., 2006(Oishi et al., , 2009. Moreover, a recent study shows that induced pluripotent stem cells (iPSCs) derived from the fibroblasts of individuals with LS display higher basal pERK levels compared with those of control iPSCs (Carvajal-Vergara et al., 2010). How the NS and LS variants of Shp2 would both activate MAPK signaling remains to be determined. Whereas catalytic activity of Shp2 is enhanced by the NS mutations, it is reduced by the LS mutations, indicating that MAPK activation is not directly affected by alterations in Shp2 catalytic activity. Rather, the NS and LS Shp2-variant-induced defects may result from a phosphatase-independent function of Shp2. Alternatively, as suggested in a recent report, the LS variants of SHP2 retain some catalytic activity and mediate gain-of-function phenotypes, because they have an increased propensity for the open conformation. Thus, the LS variants bind upstream activators preferentially and stay in complex with their scaffolding adapters longer, thus prolonging specific substrate turnover (Yu et al., 2013), which in turn may lead to MAPK activation.
Whereas MAPK activation may be elevated by both NS and LS variants of Shp2, elevated PI3K/AKT signaling is exclusively associated with LS. Hyperactivation of AKT signaling in response to LS-Shp2 is responsible for HCM in mouse hearts (Ishida et al., 2011;Marin et al., 2011;Schramm et al., 2012). We also observed elevated pAkt levels in response to LS-Shp2 in zebrafish embryos (supplementary material Fig. S3), but we did not observe HCM in these embryos. The cardiac defects in embryos expressing NS and LS Shp2 variants were indistinguishable. Moreover, these heart defects were rescued by early treatment with the MEK inhibitor to a similar extent, indicating a causal role for elevated MAPK activation in the observed cardiac defects, independently of Akt signaling.
In summary, our data provide new insights into the role of pathogenic Shp2 variants in cardiac development. Expression of NS and LS Shp2 variants led to similar cardiac defects in zebrafish that were associated with impaired L/R asymmetry, resulting from impaired cilia function in KV. Treatment with a MEK inhibitor prior to gastrulation rescued cilia function, L/R asymmetry and heart displacement, suggesting a causal relation with enhanced MAPK signaling. No studies to date have examined the effect of NS and LS Shp2 variants on L/R asymmetry. Our paper opens a new prospective in understanding how pathogenic SHP2 and downstream MAPK activation relates to cardiac development through its effects on L/R asymmetry.

Fish line
Zebrafish were maintained and the embryos were staged as previously described (Kimmel et al., 1995). The tg(myl7:GFP) line has been previously described (Chocron et al., 2007). All procedures involving experimental animals were approved by the local animal experiments committee and performed in compliance with local animal welfare laws, guidelines and policies, according to national and European law.
For zebrafish immunohistochemistry, embryos were fixed in 4% DENT's solution (20% dimethylsulfoxide/80% methanol) at 4°C, and subsequently blocked for 1 h in PBS containing 5% lamb serum, 1% BSA, 1% DMSO and 0.1% Triton-X. Embryos were incubated in mouse anti-acetylated tubulin (1:500, Sigma) for 3 h at room temperature. After extensive washing, embryos were incubated in goat anti-mouse Alexa Fluor 488. Embryos were flat-mounted in glycerol. Images were acquired using an SPE laser-scanning confocal microscope. Confocal z-series images were assembled to present the sum of the focal planes; cilia length, cilia number and KV lumen area were manually measured using ImageJ software (http://rsb.info.nih.gov/ij/).

Pharmacological inhibition of MAPK signaling
To test the prevention of the NS/LS Shp2 mRNA phenotype, embryos injected with NS/LS Shp2 mRNA were incubated with 0.25 μM CI-1040 at 4.5 hpf or 10 hpf for 1 h at 28.5°C in the dark.

Time-lapse imaging and analysis
Embryos at the 22-somite stage were dechorionated and mounted in glassbottom six-well plates using 0.25% agarose in E3 embryo medium containing 16 mg/ml 3-amino benzoic acid ethylester to block contractile movements. Confocal imaging was performed using a spinning disc confocal laser scanning microscope with 20× magnification, acquiring stacks every 5 min. Embryos were kept at 28.5°C during recordings. ImageJ software (http://rsb.info.nih.gov/ij/) was used to generate time-lapse movies and for cell counting. Cell tracking was carried out using Volocity software (Improvision) followed by manual inspection of individual tracks generating quantification of total speed (track length/time). Rotation was calculated by measuring the angle with the LR axis of four imaginary lines connecting four individual cells at the start and the end of the time-lapse (200 min). All statistical analyses were performed in Excel (Microsoft) using the two-tailed Student's t-test.

KV fluid flow and cilia motility
Fluorescent beads (Polysciences) were injected into KV to visualize flow as described previously (Essner et al., 2005). Bead flow was imaged at the 8-to 10-somite stage (SS) on a Leica AF7000 microscope (Leica Microsystems GmbH, Wetzlar, Germany) using a 63× water dipping objective and an Hamamatsu C9300-221 high-speed CCD camera (Hamamatsu Photonics, Hamamatsu City, Japan) at 80 fps to record a 10 s movie. Movies were generated using ImageJ software. Bead tracking was carried out using Volocity software (Improvision). For each embryo, four or five beads were tracked for a minimum of 50 frames of the movie and the average bead velocity was calculated.

Statistics
Cilia measurements were analyzed using the two-tailed Student's t-test. Averages for controls and experimental were compared within each clutch of embryos. Results were considered significant when P<0.05 and results are expressed as mean±s.e.m. (*P<0.05, **P<0.01, ***P<0.001).