A lmod1a mutation causes megacystis microcolon intestinal hypoperistalsis in a CRISPR/Cas9-modified zebrafish model

doi:10.21203/rs.3.rs-4847189/v1

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A lmod1a mutation causes megacystis microcolon intestinal hypoperistalsis in a CRISPR/Cas9-modified zebrafish model

https://doi.org/10.21203/rs.3.rs-4847189/v1

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Journal Publication

published 14 Aug, 2024

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Purpose

Megacystis microcolon intestinal hypoperistalsis syndrome (MMIHS) is defined as a congenital visceral myopathy with genetic mutations. However, the etiology and pathophysiology are not fully understood. We aimed to generate a gene leiomodin-1a (lmod1a) modification technique to establish a zebrafish model of MMIHS.

Methods

We targeted lmod1a in zebrafish using CRISPR/Cas9. After confirming the genotype, we measured the expression levels of the target gene and protein associated with MMIHS. A gut transit assay and spatiotemporal mapping were conducted to analyze the intestinal function.

Results

Genetic confirmation showed a 5-basepair deletion in exon 1 of lmod1a, which caused a premature stop codon. We observed significant mRNA downregulation of lmod1a, myh11, myod1 and acta2 and the protein expression of Lmod1 and Acta2 in the mutant group. A functional analysis of the lmod1a mutant zebrafish showed that its intestinal peristalsis was fewer, slower, and shorter in comparison to the wild type.

Conclusion

This study showed that targeted deletion of lmod1a in zebrafish resulted in depletion of MMIHS-related genes and proteins, resulting in intestinal hypoperistalsis. This model may have the potential to be utilized in future therapeutic approaches, such as drug discovery screening and gene repair therapy for MMIHS.

MMIHS

lmod1a

CRISPR/Cas9

zebrafish

animal model

In Japan, megacystis microcolon intestinal hypoperistalsis syndrome (MMIHS) is defined as a congenital visceral myopathy with genetic mutations. MMIHS can be strongly suspected if abdominal distension with microcolon is found and megacystis is simultaneously confirmed on ultrasonography, abdominal radiography, and a contrast study. However, a full-thickness biopsy is generally required to differentiate other allied disorders of Hirschsprung’s disease (ADHD), such as MMIHS, chronic idiopathic intestinal pseudo-obstruction (CIIP), and hypoganglionosis. A nationwide survey in Japan showed that MMIHS is an extremely rare condition with only 19 patients diagnosed to have it during the 10-year period from 2001 to 2010, which accounted for only 5.2% of all ADHD patients [1–3]. However, the consensus guidelines from the European Society for Pediatric Gastroenterology, Hepatology, and Nutrition (ESPGHAN) define pediatric intestinal pseudo-obstruction (PIPO) as a rare disabling disorder of gastrointestinal motility, characterized by symptoms of intestinal obstruction in the absence of an organic blockage and a heterogeneous and severe group of disorders, in which MMIHS is also categorized [4] Advances in genetic research have made the criteria for diagnosing MMIHS, established by ESPHAGAN, more difficult and confusing, as the monogenic mutations of actin gamma-2 (ACTG2), myosin heavy chain 11 (MYH11), myosin light chain kinase (MYLK), myosin light chain 9 (MYL9), and leiomodin-1 (LMOD1) are categorized as MMIHS [5], which means that more genetic heterogeneity related to MMIHS and possibly variation in phenotypes that can be challenging when diagnosing patients with MMIHS. This makes diagnosing patients with MMIHS more confusing where every genotype has not been identical for each phenotype which requires further study to be clearly understood. Furthermore, no radical treatment currently exists for this extremely rare and intractable disease. Current management relied on surgical interventions, such as creating jejunostomies to avoid bowel dilatation, enterocolitis after nutrition administration, total parenteral nutrition for complement loss of water and nutrient intake due to the high-output stoma, and even intestinal and/or liver transplantation, especially for patients who continue to experience malnutrition and intestinal failure-associated liver disease (IFALD) [6]

LMOD1 is a member of the LMOD protein family, with similarity to an actin capping protein called Tropomodulin. This smooth muscle cell-restricted gene is dependent on SRF/MYOCD activation [7]. A study in 2017 using a mouse model confirmed the deletion of this gene, with the model showing similar gastrointestinal and urinary bladder phenotypes to those seen in humans [8]. Another study demonstrated two compound heterozygous variants affecting Lmod1 resulting in significantly reduced protein amounts. This could be explained by disruption of the intermolecular interaction between actin and Lmod1 along with Lmod1-mediated actin nucleation impairment [8, 9]. However, the role of LMOD1 in the intestinal motility of MMIHS remains unknown.

The purpose of this study was to further investigate how a loss of lmod1a using CRISPR/Cas9-based gene modification might cause instability of the smooth muscle components, thus resulting in intestinal hypoperistalsis in zebrafish. We therefore sought to elucidate the role of Lmod1 in intestinal smooth muscle composition, and thereby help in the development of potential future therapies by creating this MMIHS zebrafish model.

Animals

Adult male and female zebrafish (RIKEN-WT) were purchased and kept at a constant temperature of 28.5°C with a light dark cycle of 14:10 h according to standard zebrafish maintenance procedures [10] To obtain eggs, adult male and female zebrafish were separated using a divider in the breeding tank overnight. The following morning, at the beginning of the light cycle, the divider was removed, and eggs were collected 5–10 minutes later. All experiments were conducted in accordance with the ethical approval of Kyushu University for animal experiments (No. A24-153-0), and genetic recombination studies (No. 4–40).

CRISPR/Cas9 mediated deletion of lmod1a gene

The CRISPR/Cas9 technique was utilized to produce deletions in exon 1 of Lmod1a gene of the zebrafish. Single-guide RNA (sgRNA) was designed using the CHOPCHOP website (https://chopchop.cbu.uib.no) and produced using the commercial EnGen® sgRNA Synthesis Kit (NEB, USA). Several sgRNAs were made, and the sgRNA with the highest deletion production efficiency was chosen. During the 1–4 cell stage of zebrafish embryos, a mixture of Cas9 protein (PNA Bio, USA) and sgRNA was microinjected at a ratio of 400:200 ng.

Genotype confirmation

At three days post-fertilization (dpf), microinjected eggs were collected for DNA extraction (DNeasy Blood and Tissue Kit, Qiagen, Germany), and PCR screening was conducted. If heteroduplexes were detected, the eggs were raised until they were 3–4 months old. Individual zebrafish fins were clipped for DNA extraction for further genotyping. A T7 endonuclease assay was conducted (EnGen Mutation Detection Kit; NEB, USA) to check for the presence of mutations in each fish. DNA was confirmed to be positive for deletion by Sanger sequencing.

Total RNA extraction and qRT-PCR

Gene expression analysis of lmod1a, myh11a, acta2 and myod1 was conducted using quantitative RT-PCR (qRT-PCR) method. Individual zebrafish whole body at 14 dpf was euthanized by overdosing the fish in 300 mg/l tricaine methane sulfate (MS222, Sigma Aldrich, USA). The tissue was then washed with PBS and continue with total RNA extraction using a commercial reagent (Monarch Total RNA Miniprep Kit, NEB, USA). Total RNA was then quantified, and purification was confirmed using the nanodrop 2000 spectrophotometer machine (ThermoFisher, USA). Approximately 100–200 ng of total RNA was then used for qRT-PCR using the Luna Universal One-Step RT-qPCR Kit (NEB, USA) according to manufacturer’s protocol in the QuantStudio 3 Real-Time PCR machine (ThermoFisher, USA). The primers used in this study can be seen in Supplementary Table 1. The relative gene expression level was calculated using the Livak method normalized with the β-actin housekeeping gene [11]

Western blotting

At approximately 3 months old, zebrafish was euthanized with the same protocol as previously mentioned. The tissue was then transferred to RIPA Buffer reagent (Nacalai Tesque, Japan) and homogenized using mechanical tissue disruptor. The samples were incubated in ice for 1 hour and centrifuged at 10.000 g for 10 minutes at 4 °C. The protein samples were then quantified using the Bradford assay method and 30 µg of protein samples was loaded in a SDS PAGE precast 5 to 20% gel. Proteins were transferred to nitrocellulose membrane and incubated with primary antibodies LMOD1 antibody (1:1000, ThermoFisher, Cat No. PA5-44224) and A-SMA antibody (1:1000, GeneTex, Cat No. GTX100034). The LMOD1 antibody we used covers the 3’ end of exon 1 of LMOD1 passing the deletion point. Secondary antibodies used were as follows: Anti-mouse IgG, HRP-linked Antibody (1:4000, Cell Signaling, Cat No. 7076) and Anti-rabbit IgG, HRP-linked antibody (1:4000, Cell Signaling, Cat No. 7074).

Intestinal peristalsis analysis

Microinjected fish were analyzed for intestinal peristalsis using the gut transit assay (GTA) method [12]At 7 dpf, both microinjected and wild type larvae were fed with fluorescent label food for 1 hour, followed by screening of the zebrafish that contained the food. The fluorescent food-containing fish were then separated. The fish were then imaged using a confocal microscope after 2, 6, 12, 18, and 24 h to check the progress of the fluorescent food. At the same time, separate fish were recorded for 5 min to observe intestinal peristalsis movement. Spatiotemporal mapping (STM) was performed using the ImageJ software program to calculate the number, frequency, interval, distance, and velocity of peristalses [12].

Statistical analysis

All variables containing numerical data were analyzed using independent t-test with SPSS (IBM, USA). P-values of less than 0.05 were considered statistically significant.

Generation of CRISPR/Cas9 mediated mutation in zebrafish

We used the zebrafish as an animal model to study the effect of the lmod1a mutation MMIHS especially the intestinal motility aspect. We targeted exon 1 of the lmod1a gene using CRISPR/Cas9-mediated deletion, this exon 1 location was chosen based on the locations of previous mutations found in human and also selected using the CHOPCHOP ranking tool (Fig. 1A). Genetic screening was first conducted using T7 Endonuclease assay showing 3 bands indicating the presence of mismatch in the DNA of zebrafish (Fig. 1B). To confirm the deletion of lmod1a gene, the DNA was then used for Sanger sequencing, which we found a 5 base-pair deletion that created a frameshift mutation in 17 amino acids (Fig. 1C). This was followed by a premature stop codon formation, which caused protein truncation (Fig. 1D).

Expression analysis

To further analyze the effect of the lmod1a deletion, we confirmed the decrease in the mRNA expression of lmod1a, myh11, myod1 and acta2 (a-sma) by qRT-PCR (Fig. 2) and the protein expression of Lmod1 and Acta2 by western blotting (Fig. 3). We found that the lmod1a mRNA expression was significantly reduced by about half along with other genes including myh11a and myod1 (Fold change: 0.47, p:0.047; 0.42, p:0.003; 0.43, p:0.001; respectively). While the mRNA expression of acta2 was the most downregulated by almost one-fifth compared to wild type (Fold change: 0.19; p:0.0001) (Table 1). Western blotting showed that both the protein expression of Lmod1 and Acta2 was reduced by almost one-fifth compared to wild type. Together, these results showed that this knockout method affected not only lmod1 but other smooth muscle related genes.

Table 1

mRNA expression of smooth muscle related genes
	Δ Cq ± SD	ΔΔCq	Fold Change	p-value	95% CI
lmod1a	13.25 ± 1.39	1.08	0.47	0.047*	0.02–2.14
myh11a	6.48 ± 0.89	1.24	0.42	0.003*	0.63–1.85
myod1	4.42 ± 0.44	1.21	0.43	0.001*	0.67–1.75
acta2	2.12 ± 0.72	2.42	0.19	0.0001*	1.69–3.13
WT	12.17 ± 0.91
*, p < 0.05; WT: wild type, Cq: cycle quantification, SD: standard deviation, CI: confidence interval

Intestinal motility functional analysis

Both GTA and STM analyses were conducted to analyze the intestinal motility function in the mutant zebrafish model. These analyses were done at 7 dpf fish due to the lack of melanin pigmentation at this time point, allowing great visibility to assess the intestine organ. The zebrafish intestine was divided into four zones for GTA analysis based on their location relative to the swim bladder (SB). Zone 1 was intestinal bulb before SB, zone 2 was located ventral to the SB, zone 3 was between the intestinal bulb and mid intestine, and zone 4 is mid and posterior intestine (Fig. 4A). The GTA showed that the mutant group (KO) had significantly higher amounts of remaining food, especially in zones 2 and 3 at 6 and 12 hours, indicating lower intestinal peristalsis (Fig. 4B and C). Furthermore, to quantify the intestinal peristalsis we did spatiotemporal mapping analysis by recording the intestinal movement for 5 minutes. The video was then analyzed with ImageJ using the Reslice tool, which then the frequency, speed, distance and interval could be calculated. The results showed that the mutant group exhibited significantly fewer peristalsis (KO vs WT, 1.21 ± 0.36 vs 1.51 ± 0.22, p: 0.04; respectively), slower (KO vs WT, 13.27 ± 2.75 vs 17.97 ± 1.78, p:<0.01; respectively), and shorter (KO vs WT, 367.03 ± 45.17 vs 526.80 ± 53.07, p: <0.01; respectively) peristalses than the wild type (WT) group (Fig. 5, Table 2).

Table 2

Statistical analysis of intestinal peristalsis spatiotemporal mapping
	Number of peristalsis	Frequency ( / min )	Interval (Δt : sec)	Distance (d : µm)	Velocity (v : µm/s)
KO (10)	5.2 ± 1.7	1.21 ± 0.36	39.23 ± 6.15	367.03 ± 45.17	13.27 ± 2.75
WT (10)	6.0 ± 0.7	1.51 ± 0.22	41.47 ± 3.61	526.80 ± 53.07	17.97 ± 1.78
p-value	0.23	0.04*	0.35	< 0.01*	< 0.01*
*, p < 0.05; KO: mutant group, WT: wild type group

MMIHS is one of the most intractable gastrointestinal disorders, occasionally detected during prenatal screening; however, most cases occur in early postnatal life. The clinical manifestations of MMIHS vary from mild to severe intestinal dysfunction, concurrent bladder dysfunction, and weak uterine constriction in adult females. This variability in disease severity indicates that visceral myopathies are multifactorial disorders involving genetic, epigenetic, and environmental modulators that are not yet well understood [13]. Due to its rarity, severity, and inherency, the Japanese government supported a nationwide survey of ADHD, including MMIHS, and revealed that the current status of pathognomonic ADHD is unestablished, and therapeutic outcome is quite unsatisfactory; 42% of patients died before 14 years of age [1, 2]. One of the reasons why MMIHS still has high lethality without radical treatment is the unclear ontogeny of MMIHS; therefore, we believe that creating a simple and easy-to-replicate animal model of MMIHS will provide new insights into the etiology and pathophysiology of the condition.

Our study demonstrated a simple and easy-to-replicate technique that induced a deletion, which reduced the expression of the lmod1a gene in zebrafish. To our knowledge, this is the first report to demonstrate zebrafish as a model for MMIHS and intestinal failure by targeting the lmod1a gene. Recently, zebrafish have been used in biomedical research, with an increasing number of papers published annually [14]. Zebrafish have several advantages over other animal models, including a fully sequenced genome (with 71.4% similarity to humans [15]), high fecundity, external fertilization, very easy genetic manipulation, rapid development, and a nearly transparent embryo [14]. Since the zebrafish intestine starts to open up at 6 dpf [16], and an intestinal function analysis can be easily accessed during this transparent larval period, we assumed that the genetically modified zebrafish model is an appropriate experimental model for MMIHS, which is an extremely rare and presumably inheritable disease.

LMOD1 is one of the three leiomodin protein isoforms that function in the actin nucleation process [7] and more recently in thin filament elongation, due to the binding of its domains (tropomodulin binding site 1 [TpmBS1], actin binding site 1 [ABS1], and actin binding site 2 [ABS2]) with the pointed end of tropomodulin [17]. Halim et al. reported that a Dutch consanguineous sibling identified as carrying a homozygous nonsense mutation of LMOD1 causing premature stop codon in exon 2, showed MMIHS with no intestinal peristalsis, hydronephrosis, bilateral distention of the ureter, distended bladder, or microcolon and died after birth [8]. They also used CRISPR/Cas-9 to induce a large deletion of 151 bp in exon one of Lmod1 in a mouse model, which caused a premature stop codon and showed phenotypes similar to MMIHS, including macrocystis, hydronephrosis, and bowel distension. Using a human smooth muscle cell line, this report also demonstrated a reduction in actin, elongated cytoskeletal dense bodies, and impaired contractility in LMOD1 knockout mice [[8]. Liu et al. demonstrated two compound heterozygous mutations of LMOD1 in a patient with PIPO (c.1106C > T, p.T369M; c.1262G > A, p.R421H), which significantly reduced the protein expression of LMOD1 and suppressed nucleation activity and polymerization rates using a cell line knockout model. They also used molecular modeling to explain the functional effect of the mutations on muscle contraction. The model showed that the intermolecular interaction of the ABS2 domain and actin was impaired [9]. However, the limitation of using mouse and cell line models prevented them from studying the hypoperistalsis effect of LMOD1 knockout due to the inability to reliably analyze intestinal peristalsis movement through gut transit analysis or other techniques in vivo [8, 9]. In this respect, our zebrafish model had an advantage over mouse models and cell lines, since both GTA and STM clearly showed significant hypoperistalsis in lmod1a knockout larvae in comparison to the wild type.

We confirmed the significant decrease in the mRNA expression of lmod1a, we also showed the lower expression of other smooth muscle-related genes, such as acta2 (a-sma), myh11, and myod1, and lower protein expression of Lmod1 and Acta2, supporting the application of this zebrafish model in the study of smooth muscle-related myopathy [18–20]. In previous research the MMIHS patient’s parents each had heterozygous mutation containing a copy of the premature stop codon and they showed no phenotype. [8] This should also be noted in our animal model, since we used F0 lineage of zebrafish which means that mosaicism can play a role in our mutations. We have confirmed through Sanger sequencing that the mutations were homozygous, however due to mosaicism some heterozygous variants can still appear in our fish, which can also explain the remaining expression of lmod1a mRNA and protein. Among the analyzed genes, the expression of acta2 was the most downregulated. The expression level was approximately five times lower than that in the control, while the expression levels of other genes were approximately two times lower. ACTA2 mutations are known to cause defects in actin filaments and actin-based spreading in both vascular and visceral smooth muscle cells [21, 22]. Although the mechanism is still unknown, the ACTA2 R179H mutation has a higher depolymerization rate compared to the WT and a 40-fold higher critical concentration for polymerization, which suggests that the severe type of MMIHS may include vascular smooth muscle myopathy [13]. MYH11 encodes the smooth muscle myosin heavy chain, a major component of the thick filaments of smooth muscle cells. MYH11 has also been reported as a target gene of MMIHS [23, 24]. A homozygous deletion of Myh11 in a mouse model also showed a distended bladder and abnormal intestinal movement, proving its loss-of-function effect on smooth muscle contractility, which may lead to MMIHS [25]. Cross-downregulation between lmoad1a and myh11 has not been reported, since further studies using next-generation sequencing or cell lines are needed to confirm this interaction. MYOD1 (myogenic differentiation 1) is a nuclear protein that regulates muscle cell differentiation (especially skeletal muscles) by inducing cell cycle arrest, which is a prerequisite for myogenic initiation. A direct relationship between MYOD1 and MMIHS has not yet been reported. Since LMOD1 plays a role in the actin nucleation process and filament elongation, we suggest that downregulation of myod1 is the subsequent effect of depolymerization and the lack of elongation of smooth muscle filaments.

The present study is associated with several limitations. First, zebrafish lack a urinary bladder; therefore, it was not possible to analyze the urinary bladder for megacystis. Second, our study was conducted in the F0 lineage of mutant fish, and we only analyzed zebrafish with confirmed deletions through Sanger sequencing, which means that any hereditary conditions of MMIHS that penetrate to the next generation was not assessed in this study. Third, our study did not clearly show compensatory pathways to preserve the remnant intestinal function in the absence of lmod1a. In the future, to complement human organ-specific smooth muscle-related phenomena and the intermolecular analysis, we need to use human smooth muscle cell lines to validate our findings and conduct next-generation sequencing to analyze the impact of lmod1a knockout on other smooth muscle-related molecules.

In conclusion, this is the first report to establish an MMIHS zebrafish model with a lmod1a mutation, which was engineered using the CRISPR/Cas9 genome editing technique. The targeted deletion of lmod1a in zebrafish resulted in intestinal hypoperistalsis, as well as the downregulation of other smooth muscle actin-binding proteins, such as Acta2. This model can be used to elucidate the mechanism of intestinal smooth muscle function and may have the potential to identify chemical substances or gene-targeting approaches for MMIHS.

CRISPR/Cas9: clustered regularly interspaced short palindromic repeats and CRISPR-associated protein 9, MMIHS: megacystis microcolon intestinal hypoperistalsis syndrome, PIPO: pediatric intestinal pseudo-obstruction, sgRNA: single-guide RNA, dpf: days post fertilization, GTA: gut transit assay, STM: spatiotemporal mapping peristalsis analysis, TpmBS: tropomodulin binding site, ABS: actin binding site.

Conflict of interest disclosure:

The authors have no conflicts of interest to disclose.

Ethical approval:

This study was approved by the ethics committee of Kyushu University for animal experiments (No. A24-153-0), and genetic recombination studies (No. 4–40).

Funding:

This work was supported by JSPS KAKENHI (Grant Number JP24K11785).

Author Contribution

K.N. and T.K. designed the study; A.S.K. and Y.T. analyzed the data; and A.S.K. and K.N. wrote the draft of the manuscript. T.S. and Z.X. contributed to data collection, cleaning, and interpretation. K.N., K.K., T.K., N.K., J.M., T.M., and T.T. critically reviewed the manuscript. The final version of the manuscript has been approved by all authors.

Acknowledgement

We would like to thank Brian Quinn for his help with the English language editing of this manuscript. We also thank Takahiro Nagasawa and Miki Nakao, Department of Bioscience and Biotechnology, Faculty of Agriculture, Kyushu University, for their assistance in this study.

Code availability:

The datasets used in this study are available from the corresponding author upon reasonable request.

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A lmod1a mutation causes megacystis microcolon intestinal hypoperistalsis in a CRISPR/Cas9-modified zebrafish model

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Abstract

Purpose

Methods

Results

Conclusion

Figures

Introduction

Methods

Animals

CRISPR/Cas9 mediated deletion of lmod1a gene

Genotype confirmation

Total RNA extraction and qRT-PCR

Western blotting

Intestinal peristalsis analysis

Statistical analysis

Results

Generation of CRISPR/Cas9 mediated mutation in zebrafish

Expression analysis

Intestinal motility functional analysis

Discussion

Abbreviations

Declarations

Conflict of interest disclosure:

Ethical approval:

Funding:

Author Contribution

Acknowledgement

Code availability:

References

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Supplementary Files

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