Upf3a but not Upf1 mediates the genetic compensation response induced by leg1 deleterious mutations in an H3K4me3-independent manner

Xie, Aixuan; Ma, Zhipeng; Wang, Jinyang; Zhang, Yuxi; Chen, Yayue; Yang, Chun; Chen, Jun; Peng, Jinrong

doi:10.1038/s41421-023-00550-2

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Published: 27 June 2023

Upf3a but not Upf1 mediates the genetic compensation response induced by leg1 deleterious mutations in an H3K4me3-independent manner

Aixuan Xie¹^na1,
Zhipeng Ma²^na1,
Jinyang Wang¹,
Yuxi Zhang²,
Yayue Chen¹,
Chun Yang ORCID: orcid.org/0000-0002-5130-8799²,
Jun Chen² &
…
Jinrong Peng¹

Cell Discovery volume 9, Article number: 63 (2023) Cite this article

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Abstract

Genetic compensation responses (GCRs) can be induced by deleterious mutations in living organisms in order to maintain genetic robustness. One type of GCRs, homology-dependent GCR (HDGCR), involves transcriptional activation of one or more homologous genes related to the mutated gene. In zebrafish, ~80% of the genetic mutants produced by gene editing technology failed to show obvious phenotypes. The HDGCR has been proposed to be one of the main reasons for this phenomenon. It is triggered by mutant mRNA bearing a premature termination codon and has been suggested to depend on components of both the nonsense mRNA-mediated degradation (NMD) pathway and the complex of proteins associated with Set1 (COMPASS). However, exactly which specific NMD factor is required for HDGCR remains disputed. Here, zebrafish leg1 deleterious mutants are adopted as a model to distinguish the role of the NMD factors Upf1 and Upf3a in HDGCR. Four single mutant lines and three double mutant lines were produced. The RNA-seq data from 71 samples and the ULI-NChIP-seq data from 8 samples were then analyzed to study the HDGCR in leg1 mutants. Our results provide strong evidence that Upf3a, but not Upf1, is essential for the HDGCR induced by nonsense mutations in leg1 genes where H3K4me3 enrichment appears not to be a prerequisite. We also show that Upf3a is responsible for correcting the expression of hundreds of genes that would otherwise be dysregulated in the leg1 deleterious mutant.

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Introduction

Living organisms have developed a variety of means to cope with environmental changes and genetic variations for their viability and fitness during evolution¹. The genetic compensation responses (GCRs), including the use of redundant genes, genetic network rewiring, alternative splicing, genetic modifiers, redundant signaling pathways, and other similar processes and mechanisms have been widely adopted and are evident across the broad range of the biological taxa². For example, regarding genetic network rewiring, by building a protein–protein interaction network containing 1870 proteins in the yeast Saccharomyces cerevisiae (S. cerevisiae), Jeong et al. found a close correlation between the lethality of a single-gene deletion with the topological position of its protein product in the web of molecular interactions³. As for genetic modifiers, through examining survival assays of ~5000 unique single-gene deletions in S. cerevisiae, Teng et al. found that most of these gene deletion strains had one additional corresponding mutant gene which appeared to be responsible for adaptive genetic changes⁴. Although the importance of the GCR for maintaining genetic robustness is generally appreciated, the field is young and the different classes of GCRs have not yet been fully fleshed out. The homology-dependent GCR (HDGCR) is an important subset of GCR, mainly due to the high abundance of homologous genes in the genome, which provides accessible resources for functional compensation. For example, the percentage of homologous genes corresponding to coding genes is approximately 65%, 63%, 72%, and 78% for the genome of Homo sapiens^5,6, Mus musculus⁷, Arabidopsis thaliana⁸ and Danio rerio (zebrafish)⁹, respectively. This neatly fits the observation that ~80% of the genetic mutants produced by gene editing technology fail to show an obvious phenotype in zebrafish¹⁰. HDGCRs induced by deleterious mutations are therefore suggested to account for this phenomenon¹¹. Recently, two reports have demonstrated that the HDGCR relies on: (1) mutant mRNA bearing a premature termination codon (PTC); (2) a homologous sequence occurring between the mutated gene and its compensatory homologs; (3) components in the nonsense mRNA-mediated degradation pathway (NMD pathway); and (4) promotion of H3K4me3 modification at the transcription start site (TSS) of the compensated genes via the complex of proteins associated with Set1 (COMPASS)^12,13. A key biological function of the NMD pathway is to cleanse the abnormal transcripts carrying a PTC¹⁴, while the COMPASS is responsible for the H3K4 methylation around the TSS site^15,16. Upf1 and Upf3b have been shown to be key positive regulators of the NMD pathway whereas Upf3a, a homolog of Upf3b, appears to play a minor or even antagonistic role in NMD pathway^14,17,18. The two previous related reports, one from our lab and the other from Stainier’s lab, have put forward two differing proposals for the factor(s) responsible for relaying the HDGCR signal from the NMD complex to the COMPASS complex. The report from Stainier’s lab proposed that Upf1 and mutant mRNA degradation are required for the activation of the HDGCR¹², whereas we showed that HDGCR depends on Upf3a¹³. Based on this discrepancy, there is a need to clarify the role of Upf1 and Upf3a in the process of the HDGCR.

Liver-enriched gene 1 (Leg1) represents a protein family highly conserved in vertebrates. It is characterized by a single domain of unknown function 781 (DUF781)^19,20,21,22. In zebrafish, Leg1 is a liver-produced novel serum protein²⁰, whereas, in the platypus and echidna, the Leg1 family member known as Monotreme Lactation Protein (MLP) is a component of milk that may function as an antimicrobial protein^23,24. The zebrafish genome contains two copies of leg1 genes, leg1a and leg1b, which are closely linked on chromosome 20²⁰. Both leg1a and leg1b transcripts are detectable at the embryonic stage, but with leg1a being the predominant form (> 90%) at this stage of development^20,21. We previously showed that knockdown using gene-specific morpholinos (MOs) of either leg1a, leg1b, or both together, resulted in a small liver at 3.5 days post-fertilization (dpf)²⁰. In contrast, except for during the winter season, knockout of the leg1a gene (the leg1a^zju1/zju1 mutant) did not affect liver development. This was likely due to an upregulation of leg1b expression discernable from 3 dpf onwards²¹, suggesting the activation of an HDGCR in the leg1a^zju1/zju1 mutant embryos.

In this report, using RNA sequencing (RNA-seq) we studied leg1b expressions in leg1a mutant embryos and distinguished the role of Upf1 and Upf3a in the HDGCR by analyzing leg1b expression in the leg1a^zju1/zju1upf3a^−/− double and leg1a^zju1/zju1upf1^−/− double mutants at either 3 dpf or 5 dpf. We also performed ultra-low-input micrococcal nuclease-based native chromatin immunoprecipitation and sequencing (ULI-NChIP-seq) analysis using the micro-dissected liver buds at 5 dpf to study the correlation between the expression of 70 liver-enriched genes and H3K4me3 modification. Our data show that Upf3a, but not Upf1, is crucial for the HDGCR in leg1a mutants at 3 dpf and 5 dpf.

Results

Only leg1a and leg1b double mutations confer a small liver phenotype

To find out whether the HDGCR is activated in the leg1a or leg1b deleterious mutants, we took the advantage of the PTC-bearing leg1a^zju1 single (with a 13 bp insertion in the exon 1 of leg1a), leg1b^zju1 single (with a 14 bp deletion in the exon 2 of leg1b) and leg1a^zju3leg1b^zju1 double (where leg1a^zju3 harbors a 7 bp deletion in leg1a) mutant lines available to our laboratory (Supplementary Fig. S1)^21,22. leg1a^zju1/zju1 single (leg1a_mu), leg1b^zju1/zju1 single (leg1b_mu) and leg1a^zju3/zju3leg1b^zju1/zju1 double (leg1_dm) homozygous mutants were all viable and fertile. Whole-mount in situ hybridization (WISH) using fabp10a as a molecular marker for the liver at 3.5 dpf showed that, compared with the wild-type controls (WT), only the leg1a^zju3/zju3leg1b^zju1/zju1 double mutant exhibited a smaller liver with neither leg1a^zju1/zju1 or leg1b^zju1/zju1 single mutants displaying such a phenotype (Fig. 1a, b). This seemed to confirm the redundant function of leg1a and leg1b in embryonic liver development²⁰. Double-probe WISH (using either fabp10a and trypsin probes together, or fabp10a and fabp2 probes together at 3.5 dpf) revealed that both the exocrine pancreas (as marked by trypsin) (Fig. 1c, d) and the intestine (as marked by fabp2) (Fig. 1e, f) exhibited a significant size reduction corresponding to that of the liver, but again this occurred only in the leg1a^zju3/zju3leg1b^zju1/zju1 double mutants. Interestingly, when comparing either the liver and exocrine pancreas (Fig. 1d, for ‘exocrine pancreas to liver ratio’) or the liver and intestine (Fig. 1f, for ‘intestine to liver ratio’) the scale of size reduction in the leg1a^zju3/zju3leg1b^zju1/zju1 double mutants was similar for both comparison, consistent with the assumption that Leg1a and Leg1b may act as a secreted signaling molecule²⁰ active in regulating the development of multiple organs/tissues²¹.

**Fig. 1: *leg1a*^zju3/zju3leg1b^zju1/zju1 double mutant exhibits a smaller liver and exocrine pancreas and thinner intestinal tube phenotype.**

The HDGCR is activated in both leg1a and leg1b single mutants

Next, we performed an RNA-seq experiment using the total RNA extracted from WT, leg1a^zju1/zju1 single (leg1a_mu), leg1b^zju1/zju1 single (leg1b_mu), and leg1a^zju3/zju3leg1b^zju1/zju1 double (leg1_dm) homozygous mutants at both 3 dpf and 5 dpf. The obtained RNA-seq data were of satisfactory quality based on the analysis of the volume of clean bases, the Clean Q30 Bases Rate, the mapping rate of the clean sequences to the zebrafish genome (Danio rerio.GRCz11) (Supplementary Tables S1, S2), and principal component analysis (PCA) (Supplementary Fig. S2a, b). We first compared the transcript levels of leg1a and leg1b and also two control genes, actin beta 2 (actb2) and betaine homocysteine S-methyltransferase (bhmt), in these four genotypes. Based on analyzing fragments per kilobase of exon model per million mapped fragments (FPKM), we found that the leg1a transcript levels were extremely low in leg1a^zju1/zju1 single and leg1a^zju3/zju3leg1b^zju1/zju1 double mutant, so did leg1b in leg1b^zju1/zju1 single and leg1a^zju3/zju3leg1b^zju1zju1 double mutants, at both 3 dpf (Fig. 2a, b) and 5 dpf (Fig. 2c, d). The results suggested that leg1a^zju1, leg1a^zju3, and leg1b^zju1 mutant mRNAs (all bearing a PTC) were subjected to degradation by the NMD pathway¹⁴. On the other hand, compared with WT controls, the levels of leg1a transcripts were approximately 50% and 44% higher in the leg1b^zju1/zju1 mutant and the levels of leg1b transcripts were 1.8- and 1.9-old higher in the leg1a^zju1/zju1 mutants at 3 dpf (Fig. 2b) and 5 dpf (Fig. 2d), respectively. No significant changes were observed for actb2 and bhmt in any of these samples. These results suggest that the HDGCR is likely activated in both leg1a^zju1/zju1 and leg1b^zju1/zju1 single mutants.

**Fig. 2: RNA-seq analysis revealed activation of GCR in both *leg1a*^zju1/zju1 single and *leg1b*^zju1/zju1 single mutants.**

To determine whether the HDGCR was indeed activated in leg1a^zju1/zju1, we compared the ratios of the leg1b precursor RNA (defined by containing intron sequences) vs total leg1b transcripts in the 36 independent RNA-seq datasets, including 9 datasets each for WT and leg1a^zju1/zju1 at both 3 dpf and 5 dpf (Supplementary Tables S1, S2, S7, S8, S13, S14), with bhmt as the control. The scenario was that the ratio of precursor RNA (un-spliced or partially spliced) could serve as an index for transcription activity²⁵. The result showed that the ratio of the leg1b precursor RNA vs total leg1b transcripts in leg1a^zju1/zju1 was significantly higher than that in the WT at 5 dpf, reaching ~267 fold that of the WT and where no significant difference was observed for the bhmt gene (Fig. 3a, right panel). This suggested activation of leg1b transcription in leg1a^zju1/zju1 mutants. Interestingly, no significant difference was observed at 3 dpf (Fig. 3a, left panel).

**Fig. 3: Knockdown of *leg1b* by MO in *leg1a*^zju1/zju1 caused a small liver phenotype.**

Next, we knocked down the Leg1b expression by injecting the morpholino (leg1b-MO) which specifically targeted the start codon ATG region of the leg1b mRNA²⁰ into the WT, leg1a^zju1/zju1 and leg1b^zju1/zju1 embryos at the one-cell stage, and then harvested the embryos at 3.5 dpf for protein extraction and WISH assay. The hypothesis here was that knockdown of the upregulated leg1b in leg1a^zju1/zju1 would lead to a small liver phenotype. Western blot assay failed to detect total Leg1 proteins in the leg1a^zju3/zju3leg1b^zju1zju1 double mutant embryos (Fig. 3b). This suggested leg1a^zju3/zju3 and leg1b^zju1/zju1 both to be null alleles. Thus, it was logical to put forward that the total Leg1 proteins detected in leg1a^zju1/zju1 represented Leg1b protein only, and vise versa, Leg1a protein only in leg1b^zju1/zju1. The leg1a^zju1/zju1 embryos contained a lower level of total Leg1 compared with WT. This is likely an outcome of the depletion of Leg1a which is the predominant form at the embryonic stage (leg1a vs leg1b FPKM, 114.9 vs 10 at 3 dpf, and 198.3 vs 14.6 at 5 dpf). The Leg1b level was further lowered by leg1b-MO injection in the leg1a^zju1/zju1 embryos (Fig. 3b). Notably, the Leg1a protein level in leg1b^zju1/zju1 was upregulated when compared with WT (Fig. 3b). Consistent with our previous report²⁰, the WISH result showed that the leg1b-MO injection had resulted in a smaller liver than that of the WT embryos. Strikingly, the leg1b-MO injection into the leg1a^zju1/zju1 embryos caused an almost liverless phenotype, which was far more severe than the leg1b-MO injected WT (Fig. 3c, d). Interestingly, injecting the leg1b-MO into the leg1b^zju1/zju1 mutant also caused a small liver phenotype. However, this remained significantly bigger than that in the WT embryos injected with leg1b-MO (Fig. 3c, d), possibly due to the binding of the leg1b-MO to the leg1b^zju1/zju1 mutant mRNA which may partially compromise the upregulation of leg1a through the HDGCR.

upf1 and upf3a transcript levels were not significantly altered in either leg1a or leg1b single mutants

To explore whether any common molecular signaling or biological pathway(s) might be mobilized to trigger the HDGCR in leg1a^zju1/zju1 and leg1b^zju1/zju1 single mutants, we carried out a further analysis of the RNA-seq data. Firstly, we checked the transcript levels of genes that had previously been shown to be involved in mediating the HDGCR (upf1, xrn1, upf3a, wdr5, rbbp5, setd1a, and ash2l)^12,13, in the RNA-seq data. No significant difference was found for any of these genes compared to WT, except ash2l (P = 0.038) in leg1a^zju1/zju1 at 3 dpf (Fig. 4a), suggesting that the transcriptional regulation of these genes might not be a prerequisite for the HDGCR activation in leg1a^zju1/zju1 and leg1b^zju1/zju1 single mutants.

**Fig. 4: *upf3a*^−/−*leg1a*^zju1/zju1 double mutant exhibits a smaller liver phenotype.**

Next, we analyzed differentially expressed genes (DEGs) by focusing on the RNA-seq data obtained from the larvae at 3 dpf (as the HDGCR had been observed at this time point) (Fig. 2). 1284 downregulated and 570 upregulated DEGs in leg1a^zju1/zju1 and 696 downregulated and 659 upregulated DEGs in leg1b^zju1/zju1 were identified (Supplementary Tables S3, S4). leg1a^zju1/zju1 and leg1b^zju1/zju1 shared 334 downregulated and 182 upregulated DEGs (Supplementary Fig. S2c). We then undertook gene ontology (GO) analysis of the 334 shared downregulated DEGs. Those significantly affected processes under the terms of molecular function (MF) and biologic process (BP) were mainly related to metabolic activities, while genes with a product related to the extracellular space and region were found under the cellular component term (CC) (Supplementary Fig. S2d). Analysis of the 182 shared upregulated DEGs identified a few metabolic pathways under the MF and BP terms, however, with only a handful of genes (Supplementary Fig. S2e). To gain more insight into the 182 shared upregulated DEGs, we manually searched the known or putative functions of these DEGs in the Zebrafish Information Network (ZFIN) (https://zfin.org/). We noticed that 23 of these DEGs were related to DNA-binding and transcription regulation (Supplementary Fig. S2f). However, whether the upregulation of these 23 DEGs was a reason for, or a consequence of, the HDGCR, or due to the loss-of-function of Leg1a/Leg1b remains unknown. This is primarily due to a number of factors: (1) the expression of leg1a and leg1b is relatively liver-specific, so the HDGCR induced by leg1a^zju1/zju1 or leg1b^zju1/zju1 mutant mRNA is presumably restricted to within the liver cells; (2) while the RNA samples were prepared from the whole larvae at 3 dpf, when the liver volume vs that of the whole larvae is less than 0.1%²⁶, detection of the HDGCR-related gene transcripts may be obscured; and (3) Leg1a and Leg1b are secretory proteins whose function might not be limited to the liver, and thus the loss-of-function of leg1a or leg1b might affect gene expression profiles in multiple organs/tissues.

To gain an insight into the biological function of Leg1 in zebrafish, we then compared the RNA-seq data from the WT and leg1a^zju3/zju3leg1b^zju1/zju1 double mutant samples. DEseq2 analysis identified 904 DEGs (378 upregulated and 526 downregulated) at 3 dpf and 875 DEGs (453 upregulated and 422 downregulated) at 5dpf (leg1_dm vs WT:|log₂(fold-change) |≥ 1, p_{ad j} < 0.05) (Supplementary Tables S3–S6). GO analysis revealed the ‘intracellular signal transduction’ category under the ‘BP’ term to be the process most significantly affected in the leg1a^zju3/zju3leg1b^zju1/zju1 double mutant for both downregulated and upregulated DEGs at both 3 dpf and 5 dpf (Supplementary Fig. S3a, b). The ‘intracellular signal transduction’ process contained 27 genes at 3 dpf and 34 genes at 5d pf of the downregulated DEGs (3 dpf and 5 dpf sharing 21 genes) (Supplementary Fig. S3c) and 24 genes at 3 dpf and 26 genes at 5 dpf of the upregulated DEGs (3 dpf and 5 dpf sharing 15 genes) (Supplementary Fig. S3d). Surprisingly, many of these genes had not been genuinely annotated and were only assigned with ENSEMBL gene identity numbers (https://asia.ensembl.org/index.html). Based on cross-species homology analysis, we noticed that a large proportion of the above genes, regardless of whether downregulated or upregulated, were predicted to encode proteins with putative ATP-binding activity and show homology to the NOD-like receptor pyrin domain-containing (Nlrp) family proteins (Supplementary Fig. S3c, d). Nlrp proteins play important roles in mediating the innate immune response upon pathogen infections^27,28,29. GO analysis, therefore, suggested that Leg1 might act as a secreted signaling molecule active in regulating the innate immune response and other biological processes.

upf3a and leg1a double mutations cause a small liver phenotype

To determine the role of upf3a and upf1 in the HDGCR in leg1a^zju1/zju1, we obtained upf3a^−/−leg1a^zju1/zju1 (upf3a;leg1a_dm) and upf1^−/−leg1a^zju1/zju1 (upf1;leg1a_dm) double mutants, respectively (Supplementary Fig. S4a, b). The upf3a^−/−leg1a^zju1/zju1 double mutant was viable and fertile, just as were the upf3a^−/− single (upf3a_mu) and leg1a^zju1/zju1 single (leg1a_mu) mutants^13,21. However, the upf1^−/−leg1a^zju1/zju1 double mutant showed embryonic lethality, just as did the upf1^−/− single (upf1_mu) mutant¹³. Consistent with our previous report¹³, upf3a^−/− mutants displayed a slightly larger liver than that in the WT at 3.5 dpf as revealed by WISH using an fabp10a probe (Fig. 4b, c). Interestingly, the maternal-zygotic upf3a^−/−leg1a^zju1/zju1 embryos developed small livers, resembling the leg1a^zju3/zju3leg1b^zju1zju1 double mutant (Fig. 4b, c). Considering the fact that the leg1a^zju1/zju1 mutant developed a normal-sized liver and the upf3a^−/− mutant with a slightly enlarged liver, the small liver phenotype displayed by the upf3a^−/−leg1a^zju1/zju1 double mutant is proposed to be the outcome of a compromised HDGCR leading to a reduction in leg1b expression in the upf3a^−/−leg1a^zju1/zju1 double mutant. This hypothesis was supported by the RNA-seq data analysis (Fig. 5a–d) and by our western blot assay which showed that, compared with the leg1a^zju1/zju1 single mutant, the Leg1b protein level was reduced in the upf3a^−/−leg1a^zju1/zju1 double mutant (Fig. 4d). However, we cannot currently exclude the possibility that a concomitant loss-of-function of upf3a and leg1a might had an accumulative effect to cause the observed small liver phenotype. The upf1^−/−leg1a^zju1/zju1 double mutant, resembling the upf1^−/− single mutant¹³, exhibited smaller livers compared to WT(Fig. 4e, f). This suggests that the upf1 deleterious mutation is genetically epistatic to leg1a^zju1/zju1, despite the continued upregulation of leg1b expression in the upf1^−/−leg1a^zju1/zju1 double mutant (Fig. 5e–h).

**Fig. 5: RNA-seq analysis showed that the upregulated *leg1b* transcript level in *leg1a*^zju1/zju1 was downregulated by Upf3a depletion but was unaffected by Upf1 depletion.**

Depletion of Upf3a but not Upf1 abolishes the upregulation of leg1b caused by the leg1a mutation

Total RNA extracted from WT, leg1a^zju1/zju1 single (leg1a_mu), upf3a^−/− single (upf3a_mu) and upf3a^−/−leg1a^zju1/zju1 double mutant (upf3a;leg1_dm) at 3 dpf and 5 dpf (three independent samples for each genotype) was subjected to RNA-seq analysis (Supplementary Fig. S5a, b and Tables S7–S12). PCA analysis revealed that, except for one upf3a^−/− sample at 5 dpf, three independent samples for each genotype were neatly clustered (Supplementary Fig. S5a, b). The one non-clustered sample was excluded from further analysis. Expression analysis based on FPKM revealed that the leg1a mutant mRNA was at a very low level, not only in the six leg1a^zju1/zju1 single mutant samples, but also in the six upf3a^−/−leg1a^zju1/zju1 double mutant samples when compared to the WT samples at 3 dpf and 5 dpf. No significant changes were found relating to the two control genes bhmt and actb2 (Fig. 5a–d). This suggested that the leg1a mutant mRNA had been subjected to degradation by an active NMD pathway in these genetic backgrounds, and thus confirmed Upf3a not to be a positive regulator of the NMD pathway¹⁷. In contrast, the leg1b mRNA levels were upregulated in all six leg1a^zju1/zju1 single mutant samples as compared to the WT samples at both 3 dpf and 5 dpf (Fig. 5a–d). However, the upregulated leg1b expression was significantly downregulated in the six upf3a^−/−leg1a^zju1/zju1 double mutant samples as compared to the leg1a^zju1/zju1 single mutant and was returned to a similar level of that in the WT at both 3 dpf and 5 dpf (Fig. 5a–d). This suggested that Upf3a depletion had compromised the HDGCR in leg1a^zju1/zju1.

Three independent RNA samples from WT, leg1a^zju1/zju1 single (leg1a_mu), upf1^−/− single (upf1_mu) and upf1^−/−leg1a^zju1/zju1double (upf1;leg1a_dm) mutant at 3 dpf and 5 dpf were also subjected to RNA-seq analysis (Supplementary Fig. S5c, d and Tables S13–S18). We again observed a very low level of leg1a mutant mRNA in the six leg1a^zju1/zju1 single mutant samples at 3 dpf and 5 dpf (Fig. 5e–h). In contrast, the levels of leg1a mutant mRNA were significantly higher in the upf1^−/−leg1a^zju1/zju1 double mutant than in the leg1a^zju1/zju1 single mutant (Fig. 5e–h), likely due to the inactivation of the NMD pathway after depleting Upf1 in the leg1a^zju1/zju1 background¹⁴. Expression analysis also revealed the upregulation of the leg1b expression in all six leg1a^zju1/zju1 single mutant samples compared to the WT samples at 3 dpf and 5 dpf (Fig. 5e–h). Notably, the leg1b mRNA levels in the upf1^−/−leg1a^zju1/zju1 double mutant were not significantly different from those in the leg1a^zju1/zju1 single mutant (Fig. 5e–h), suggesting that Upf1 does not play an obvious role in the activation of the HDGCR in leg1a^zju1/zju1.

Strikingly, further hierarchical analysis of the 366 upregulated and 718 downregulated DEGs at 3 dpf, and 473 upregulated and 746 downregulated DEGs at 5 dpf between the leg1a^zju1/zju1 mutant and WT identified by the RNA-seq experiment (Supplementary Tables S9–S12), showed that 263 (3 dpf) and 327 (5 dpf) upregulated and 493 (3 dpf) and 520 (5 dpf) downregulated DEGs were obviously deregulated in the upf3a^−/−leg1a^zju1/zju1 double mutant. In contrast, the expression patterns of the majority of the DEGs identified in the leg1a^zju1/zju1 single mutant were not drastically altered in the upf1^−/−leg1a^zju1/zju1 double mutant (Fig. 6a–d). These data suggest that depleting Upf3a in leg1a^zju1/zju1 had blocked the HDGCR and altered gene expression profiles. Interestingly, GO analysis of the DEGs altered by the Upf3a depletion at 3 dpf (Supplementary Tables S9, S10) showed that some metabolic pathways under the GO_MF and GO_BP terms and ‘membrane’ and ‘integral component of membrane’ categories under the GO_CC term were significantly affected for the altered downregulated DEGs (Supplementary Fig. S5e). However, only the ‘extracellular space’ category under the GO_CC term was outstanding for the altered upregulated DEGs (Supplementary Fig. S5f). At 5 dpf, ‘RNA polymerase II transcription factor activity’ and ‘RNA polymerase II regulatory region’ were two categories identified for both the downregulated and upregulated DEGs altered by Upf3a depletion (Supplementary Fig. S5g, h) but each constituting of distinct genes (Supplementary Tables S11, S12). Meanwhile, the ‘intracellular signal transduction’ category for the altered downregulated DEGs and the ‘extracellular space/region’ and ‘mitochondrion’ categories for the altered upregulated DEGs at 5 dpf (Supplementary Tables S11, S12) were also identified by the GO analysis (Supplementary Fig. S5g, h). However, how and whether these DEGs were affected by the Upf3a depletion remains unknown.

**Fig. 6: Depletion of Upf3a but not of Upf1 obviously alters the DEGs in the *leg1a*^zju1/zju1 mutant.**

It is intriguing that upf3a^−/− and upf1^−/− had distinct effect upon DEGs in leg1a^zju1/zju1 where both upf3a^−/−leg1a^zju1/zju1 and upf1^−/−leg1a^zju1/zju1 double mutants displayed a small liver phenotype (Fig. 4b, c, e, f). This implies that distinct liver developmental genes might be altered by upf3a^−/− and upf1^−/−. Zebrafish liver development is governed by a genetic network formed by genes encoding multiple signaling molecules (e.g., Bmp, Fgf and Wnt, etc.) and transcription factors (such as Foxa and Gata family members)^30,31,32,33. When compared with the WT, analysis of the RNA-seq data identified 647 upregulated and 933 downregulated DEGs in upf3a^−/−leg1a^zju1/zju1 double mutant, and 1264 upregulated and 766 downregulated in upf1^−/−leg1a^zju1/zju1 double mutant at 3 dpf (Supplementary Tables S9, S10, S15, S16). Cross-comparison showed that upf3a^−/−leg1a^zju1/zju1 and upf1^−/−leg1a^zju1/zju1 double mutants shared 74 upregulated and 121 downregulated DEGs, respectively. Despite this, no well-known genes controlling liver development were among these shared genes (Supplementary Fig. S6a). We then examined the distinct downregulated DEGs and found that genes related to the BMP signaling (bmp2a, bmp3, and bmp7b) and Fgf signaling (fgf12a) were significantly downregulated in upf1^−/−leg1a^zju1/zju1 double mutant whereas signaling molecules fgf1b and fgf20b and transcription factors hnf1a, gata1a, and pparaa were identified in upf3a^−/−leg1a^zju1/zju1 double mutant (Supplementary Fig. S6b, c).

To discover whether Upf3a is also required for the compensatory expression of leg1a in the leg1b^zju1/zju1 mutant, we injected upf1 and upf3a specific MOs¹³ (Supplementary Fig. S7) into the fertilized WT and leg1b^zju1/zju1 eggs at the one-cell stage and harvested the embryos at 1.5 dpf for total RNA extraction. Quantitative rea-time PCR (qPCR) analysis of the leg1a transcripts showed that upf3a-specific MO but not the upf1-specific MO significantly downregulated the leg1a expression in the leg1b^zju1/zju1 mutant (Fig. 6e).

Upregulation of leg1b caused by the leg1a mutation is not coupled with H3K4me3 modification

Upf3a is proposed to interact with Wdr5 (a COMPASS component) to mediate the HDGCR through enhancing H3K4me3 in the promoter region of compensatory genes¹³. We, therefore, micro-dissected the liver buds from WT, leg1a^zju1/zju1 single (leg1a_mu), upf3a^−/− single (upf3a_mu), and upf3a^−/−leg1a^zju1/zju1 double (upf3a;leg1a_dm) mutant embryos at 5 dpf (Supplementary Fig. S8a) for ULI-NChIP-seq analysis of the distribution of H3K4me3 on chromatin (Supplementary Table S19). Normalized reads distribution profiles showed that H3K4me3 was enriched around the transcription start sites (TSS) of 11,197 genes (Fig. 7a; Supplementary Table S20), including housekeeping genes actb2, pck1 and gapdh and liver-enriched cpn1, rgrb, gstp1 and tpt1 in all four genotypes (Fig. 7b; Supplementary Fig. S8b, c). As expected, H3K4me3 was not enriched around the TSS sites of the muscle gene myl7, myog, myod1 and myoz2b or the intestine gene chia.1 and alpi.2 in the liver cells (Fig. 7c, d; Supplementary Fig. S8d). Surprisingly, ULI-NChIP-seq results did not reveal any obvious changes in the level of H3K4me3 around the TSS region of leg1b among all four genotypes. There was also no specific TSS-enrichment of H3K4me3 for either the leg1a or leg1b gene (Fig. 7e).

**Fig. 7: Upregulation of *leg1b* in *leg1a*^zju1/zju1 is not coupled with an enrichment of H3K4me3 around its TSS.**

To investigate whether H3K4me3 modification around the TSS region is a hallmark for the expression of liver-enriched genes, we compared the patterns of H3K4me3 modification between the adult and embryonic liver on the genomic regions of the 70 genes (the expression of which had been experimentally proved to be enriched in both adult and embryonic liver and are therefore termed as liver-enriched genes, including leg1a and leg1b) (Supplementary Table S21)¹⁹. The zebrafish adult liver ChIP-seq data was extracted from the available public database³⁴. The result showed that among the 47 liver-enriched genes showing an obvious TSS-enrichment of H3K4me3 in the adult liver only 27 (including agt and cnp) displayed enrichment (Fig. 8a; Supplementary Fig. S9a), while another 20 (including leg1a, leg1b, cp, and gatm) lacked any obvious TSS-enrichment in the embryonic liver (Figs. 7e and 8b; Supplementary Fig. S9b). For the 22 liver-enriched genes displaying an H3K4me3 enrichment across the TSS and along the adjacent gene body region in the adult liver, 13 genes (including c9 and nupr1) showed similar H3K4me3 enrichment patterns (Fig. 8c; Supplementary Fig. S9c) while the remaining 9 genes (inclduing fga and uox) were drastically different in the embryonic liver (Fig. 8d; Supplementary Fig. S9d). For one gene, zar1, no H3K4me3 enrichment was detected in the adult liver, whilst such enrichment was observed in the embryonic liver (Supplementary Fig. S9e).

**Fig. 8: Cross-comparison of H3K4me3 enrichment in the genomic regions of 70 liver-enriched genes between WT adult and embryonic liver.**

Discussion

In this report, we have shown that leg1b is robustly upregulated in the leg1a^zju1/zju1 mutant. This clearly suggested that the activation of the HDGCR had been triggered by the leg1a deleterious mutation. RNA-seq analysis of the upf3a^−/−leg1a^zju1/zju1 and upf1^−/−leg1a^zju1/zju1 double mutants implicated that Upf3a, but not Upf1, was necessary for the HDGCR triggered by the leg1a^zju1/zju1 deleterious mutation. This role of Upf3a was supported by the small liver phenotype displayed by the upf3a^−/−leg1a^zju1/zju1 double mutant, mimicking the liver phenotype observed in the leg1a^zju3/zju3leg1b^zju1/zju1 double mutant. Interestingly, the DEGs between leg1a^zju1/zju1 and WT partially reverted to a WT-like pattern in the upf3a^−/−leg1a^zju1/zju1 double mutant, whereas the DEGs showed a similar pattern between the leg1a^zju1/zju1 single and upf1^−/−leg1a^zju1/zju1 double mutants, further suggesting the important role of Upf3a in the HDGCR. However, whether Upf3a is a general mediator of the HDGCR or is only specific to a limited class of genes, requires further validation via extensive genetic and molecular analysis. Meanwhile, we cannot exclude the possibility that Upf1 may be involved in mediating the HDGCR for a distinct group of genes. Furthermore, although the data from leg1 mutants is consistent with the induction of the HDGCR, we do not know if it is specifically induced by PTC-bearing mRNA and we cannot rule out the possibility that another class of GCRs could be responsible.

One intriguing question relates to the reason for the difference in the role of Upf3a and Upf1 in the HDGCR process. Our sequencing results provided evidence that Upf3a is not a positive regulator of the NMD pathway. This is consistent with the previous report of Shum et al. implicating mammalian Upf3a as an inhibitor, rather than a positive regulator, of this pathway, probably via competing with Upf3b in the interaction with Upf2¹⁷. Based on Shum et al.’s hypothesis, we previously proposed that both Upf3a and Upf3b could interact with the complex of PTC-bearing mRNA and the exon junction complex (EJC). When Upf3b forms a complex with Upf2, this complex will be recruited by Upf1 to mediate PTC-bearing mRNA degradation. However, if Upf3a forms the complex with Upf2, it may guide the PTC-bearing mRNA to promote the transcription of compensatory genes¹³. To understand how Upf3a mediates such an HDGCR signal, a future study may be necessary to further identify the Upf3a protein interactomes. Meanwhile, more concrete evidence is required to define the reason for the difference in the role of Upf3a and Upf1 in the HDGCR process. We speculate that the position of the PTC in the mutant mRNA or different auxiliary factors (protein or RNA) associated with the EJC may be important in determining the choice.

H3K4me3 modification at the TSS is often connected to gene activation³⁴. Through analyzing the ULI-NChIP-seq data we found that the activation of leg1b in the leg1a^zju1/zju1 mutant was not coupled with an increase in H3K4me3 modification around the TSS. This finding is discrete from the previous two reports showing an increase of H3K4me3 enrichment around the TSS of the compensatory genes^12,13. This may suggest that there could be different ways to activate compensatory genes during HDGCR. Cross-comparing the H3K4me3 counts for 70 known liver-enriched genes between the adult and embryonic liver revealed that proportions of these genes are not coupled with an enrichment of H3K4me3 around the TSS, and this phenomenon is even more widespread in the embryonic liver. These observations suggest that, in addition to the H3K4me3 modification around the TSS-site, there might be other transcriptional regulators involved in regulating embryonic liver-enriched genes. 23 genes encoding DNA-binding proteins or transcription regulators were found to be upregulated in the leg1a^zju1/zju1 mutant. These genes might serve as candidates for future study for their possible roles in upregulating the expression of leg1b in the leg1a^zju1/zju1 mutant. Alternatively, other epigenetic modifications could be examined around the leg1b locus. However, based on the low FPKM for leg1b (~14.6 for leg1b vs 198.3 for leg1a at 5 dpf), we cannot exclude the possibility that the enrichment of H3K4me3 around the leg1b TSS may not be discernable by the current ULI-NChIP-seq analysis.

Genomes usually contain many gene families and the members of each family share different degrees of homology^5,6,9,35,36. Whether fully or partially, living organisms often take the advantage of redundant functions of homologous genes to cope with genetic mutations and to maintain genetic robustness^2,11. This clearly represents an important mechanism of GCR. The HDGCR has been applied to explain the observation that ~80% of genetic mutations generated by gene editing approaches do not show an obvious phenotype in zebrafish^11,37. It is envisaged that homologous genes may co-express within the same cell so that they can compensate for each other’s function when one is deleteriously mutated. However, many homologous genes are differentially expressed in different cell types and the expression of a homologous gene (or genes) has to be activated when the predominant gene is deleteriously mutated, such as was the case for leg1a and leg1b in this study²¹. The mechanism as to how the compensatory gene is activated remains to be elucidated. Our main finding is that the upregulation of leg1b in the leg1a^zju1/zju1 mutant is dependent on Upf3a, but not Upf1, and is not coupled with an increase in H3k4me3 around the TSS. This strongly suggests that there might be a complex network or various pathways in either the triggering or regulation of HDGCRs.

Materials and methods

Zebrafish lines and maintenance

The zebrafish (Danio rerio) AB strain was used as WT in this study. The leg1a^zju1 (with 13 bp insertion in the exon 1 of leg1a) single, leg1b^zju1 (with 14 bp deletion in the exon 2 of leg1b) single, and leg1a^zju3leg1b^zju1 double (leg1a^zju3 harboring a 7 bp deletion in leg1a) mutant lines were in the AB background and were generated as previously described^21,22. The upf3a^−/− and upf1^−/− mutant lines were in the Tübingen background and were generated as described¹³. Fish were raised and maintained according to the standard procedures described in ZFIN (http://www.zfin.org).

Sampling of experiments

More than 5 pairs of parental male and female fish (WT or mutant) were used to set one specific cross. For a cross between homozygous parents, more than 23 embryos were used. For a cross between heterozygous parents, more than 100 embryos were obtained and were genotyped using gene-specific primers (Supplementary Table S22) to obtain the corresponding homozygous mutant embryos as previously described^13,22. All sets of experiments were repeated at least three times except the ULI-NChIP-seq which was performed with two repeats.

WISH and organ size measurement

WISH was performed as described previously³⁸. Probes were labeled with digoxigenin (DIG). Plasmids containing fabp10a, trypsin, and fabp2 gene fragments were obtained and reported previously³⁸. The sizes of the liver (marked by the fabp10a probe), exocrine pancreas (marked by the trypsin probe), or intestinal tube (by the fabp2 probe), were measured as previously described²⁰. In brief, after WISH, liver, exocrine pancrea, or intestinal tube was marked out using their marker genes and then imaged by Nikon AZ100 from left lateral view, after aligning the two eyes of the embryo vertically, or from a dorsal view. The pixels of the mark gene staining of the positive area in each image, as cumulated by the Adobe Photoshop CC 2018 software, were then used as the index of the liver, exocrine pancreas, or intestine sizes, respectively.

MO injection

MOs were purchased from Gene Tools (Philomath, USA). The upf3a-MO, upf1-MO, leg1a-MO, and leg1b-MO were designed as described in previous studies^13,20. A human β-globin (HBB) antisense morpholino was used as the standard control (st-MO) for upf-MO injection. A 5-base mismatch morpholino (5’-CCATgTCAcACATgTAGCAcGAgTG-3’) was designed as the mismatch control (mismatch-MO) for leg1-MO injection. To investigate the mechanism of GCR in leg1b^zju1, 1 nL of upf3a-MO (0.25 nmol/µL), upf1-MO (0.25 nmol/µL), or 1 nL st-MO control (0.25 nmol/µL) was injected into one-cell stage embryos. To investigate the function of leg1a or leg1b in leg1b^zju1 or leg1a^zju1, 1 nL leg1a-MO (0.25 nmol/µL), leg1b-MO (0.25 nmol/µL) or mis-MO was injected into one-cell stage embryos. To test the efficiency of upf1-MO and upf3a-MO, 1 nL pcs2 + 5’ UTR:GFP plasmid (80 ng/µL) or 1 nL upf3a-MO (0.25 nmol/µL) with 5’ UTR:GFP plasmid (80 ng/µL) or 1 nL upf1-MO (0.25 nmol/µL) with 5’ UTR:GFP plasmid (80 ng/µL) was injected into one-cell stage embryos. After 1 day post injection, embryos were collected and imaged using a fluorescence microscope (KEYENCE BZ-X800). The sequences of all MOs are provided in Supplementary Table 22.

Western blot

Total protein was extracted using an extraction buffer (63 mM Tris-HCl, pH 6.8, 10% glycerol, 5% β-Mercaptoethanol, 3.5% SDS) containing 1× Complete (Roche, 11873580001). Western blot was performed as described previously²⁰, using monoclonal antibody against zebrafish Leg1 as the primary antibody. Beta-Tubulin antibody (Cat. #AC021) was purchased from ABclonal company (Wuhan, China).

qPCR

qPCR was performed as described previously²⁰. Total RNA was extracted from WT or leg1b^zju1 embryos injected with st-MO, upf3a-MO, or upf1-MO at 1.5 dpf using a TRIZOL reagent (AidLab). For qPCR, total RNA was treated with DNaseI before reverse transcription and purified using an RNA clean kit (AidLab). qPCR was performed in a CFX96TM Real-Time System (Bio-Rad) using a C1000 Thermal Cycle (Bio-Rad) according to the manufacturer’s instructions. Total RNA was normalized to the zebrafish actb1 gene. The primer pairs used are listed in Supplementary Table 22.

RNA-seq analysis

As stated in the text, total RNA was extracted from the embryos of specific genotypes using a TRIZOL reagent (AidLab) according to the manufacturer’s protocol. Three independent RNA samples were obtained for each genotype in each set of RNA-seq analysis. Isolation of mRNA, library construction, high throughput sequencing, and data filtering to obtain clean reads, were performed by Novogene company (Beijing, China). Clean reads were mapped to the zebrafish genome (Danio_rerio.GRCz11) with Hisat2 (v 2.2.1)³⁹. FPKM was obtained for each gene in the RNA-seq data using the feature Counts (v 2.0.1) package⁴⁰. DEGs were obtained using DESeq2 R package (v 1.30.1)⁴¹.

ULI-NChIP-seq

ChIP was performed using the ULI-NChIP protocol as described previously⁴² with 150 livers micro-dissected from the WT, leg1a^zju1/zju1 single, upf3a^−/− single, and upf3a^−/−leg1a^zju1/zju1 double mutant embryos at 5 dpf. Two independent ChIP samples were obtained for each genotype in each set of ULI-NChIP-seq analyses. After washing with cold PBS, liver cells were lysed in cell lysis buffer (10 mM Tris-HCl, pH 7.5, 10 mM NaCl, 0.5% NP-40, 1× EDTA-free protease inhibitor cocktail, and 1 mM phenylmethanesulfonyl fluoride (Sigma) on ice for 10 min. The nuclei were pelleted at 3500 rpm for 5 min at 4 °C, and then re-suspended in nuclear extraction buffer (10 mM Tris-HCl, pH 8.0, 140 mM NaCl, 5 mM MgCl, 0.6% NP-40, 1× EDTA-free protease inhibitor cocktail and 1 mM phenylmethanesulfonyl fluoride on ice for 15 min. Chromatin was fragmented by using MNase (NEB, Cat. #M0247S) (final concentration 4U/µL) for 5 min at 37 °C. The reaction was stopped by adding 16.7 mM EDTA, 8.4 mM EGTA, 1% Triton X-100, and 1% sodium deoxycholate. Fragmented chromatin was diluted in Complete immunoprecipitation buffer (20 mM Tris-HCl, pH 8.0, 2 mM EDTA, 150 mM NaCl, 0.1% Triton X-100, 1× EDTA-free protease inhibitor cocktail and 1 mM phenylmethanesulfonyl fluoride). Chromatin was pre-cleared with 5 µL of 1:1 Protein A + G Magnetic Beads (Sigma, Cat. #16-663) and IPed with 2 μg of H3K4me3 antibody (Abcam, Cat. #ab8580)-beads complexes overnight at 4 °C with rotation. IPed complexes were washed twice with 150 µL of low salt wash buffer (20 mM Tris-HCl, pH 8.0, 2 mM EDTA, 150 mM NaCl, 1% Triton X-100, 0.1% SDS, 1× EDTA-free protease inhibitor cocktail and 1 mM phenylmethanesulfonyl fluoride) and twice with 150 µL of high salt wash buffer (20 mM Tris-HCl, pH 8.0, 2 mM EDTA, 500 mM NaCl, 1% Triton X-100, 0.1% SDS, 1× EDTA-free protease inhibitor cocktail and 1 mM phenylmethanesulfonyl fluoride). The bead-bound DNA was then eluted using ChIP elution buffer (100 mM NaHCO₃ and 1% SDS) for 3 h at 68 °C. After treatment with proteinase K and RNase A, eluted chromatin was purified using phenol-chloroform extraction, and raw ChIP material was re-suspended in 10 mM Tris-HCl, at pH 8.0. Library construction, high throughput sequencing, and data filtering to get the clean reads were performed by the Anoroad company. Clean reads were mapped to the zebrafish genome (Danio_rerio.GRCz11) using the BWA (v 0.7.17)⁴³. Regions of H3K4me3 enrichment over background were identified by using the MACS2 (v 2.2.7.1) peak calling software⁴⁴. Normalized read distribution profiles were obtained using Deeptools (v 3.2.1)⁴⁵.

Statistical analysis

Student’s t-tests were used for statistical comparisons (*P < 0.05; **P < 0.01; ***P < 0.001; ns, no significant difference).

Data availability

All relevant data are available from the authors and/or included in the manuscript or supplementary information. RNA-seq data and ULI-NChIP-seq data in this paper have been deposited in the NCBI database (BioProject: PRJNA874604).

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Acknowledgements

We thank all other members from J.P. and J.C. labs for their valuable suggestions and technical help. We thank Dr. Li Jan Lo and Ms. Xiangfeng Shen for their help in the fish facility. This work is financially supported by the National Natural Science Foundation of China (31830113, 31571495) and the National Key R&D Program of China (2018YFA0800500).

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These authors contributed equally: Aixuan Xie, Zhipeng Ma.

Authors and Affiliations

1MOE Key Laboratory of Biosystems Homeostasis & Protection, College of Animal Sciences, Zhejiang University, Hangzhou, Zhejiang, China
Aixuan Xie, Jinyang Wang, Yayue Chen & Jinrong Peng
College of Life Sciences, Zhejiang University, Hangzhou, Zhejiang, China
Zhipeng Ma, Yuxi Zhang, Chun Yang & Jun Chen

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Aixuan Xie
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Jinyang Wang
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Contributions

J.P., Z.M., and J.C. conceived the project and designed the experiments. A.X. prepared RNA samples for RNA-seq and DNA for ULI-NChIP-seq experiments and performed WISH. Z.M. generated the upf3a^−/− and upf1^−/− mutants. J.W. generated the leg1b^−/− mutant and helped in genotyping, WISH, and RNA sample preparation. Y.Z., Y.C., and C.Y. helped with the analysis of the RNA-seq and ULI-NChIP-seq data. J.P., A.X., Z.M., and J.C. wrote the manuscript.

Corresponding authors

Correspondence to Zhipeng Ma, Jun Chen or Jinrong Peng.

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Supplementary information

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Xie, A., Ma, Z., Wang, J. et al. Upf3a but not Upf1 mediates the genetic compensation response induced by leg1 deleterious mutations in an H3K4me3-independent manner. Cell Discov 9, 63 (2023). https://doi.org/10.1038/s41421-023-00550-2

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Received: 24 September 2022
Accepted: 29 March 2023
Published: 27 June 2023
DOI: https://doi.org/10.1038/s41421-023-00550-2