Nrf2 activation attenuates genetic endoplasmic reticulum stress induced by a mutation in the phosphomannomutase 2 gene in zebrafish

Edited by Igor B. Dawid, The Eunice Kennedy Shriver National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, and approved January 31, 2018 (received for review August 9, 2017)
February 22, 2018
115 (11) 2758-2763

Significance

Nrf2 is a master regulator of the antioxidant response and xenobiotic metabolism. In this paper, we demonstrate that Nrf2 also plays a critical role in the endoplasmic reticulum (ER) stress response using a zebrafish mutant in which Nrf2 is spontaneously activated. The gene responsible for this mutant was phosphomannomutase 2 (pmm2), the enzyme required for the N-glycosylation. Human PMM2 is known to be the gene responsible for PMM2-CDG (congenital disorders of glycosylation), which currently has no therapeutic options. pmm2 mutant larvae showed up-regulated ER stress and ER stress-dependent Nrf2 activation. Of note, the ER stress in mutant larvae was attenuated following treatment with the Nrf2 activator sulforaphane, suggesting that the pathological conditions of ER stress-associated diseases may be improved by taking Nrf2-activating foods.

Abstract

Nrf2 plays critical roles in animals’ defense against electrophiles and oxidative stress by orchestrating the induction of cytoprotective genes. We previously isolated the zebrafish mutant it768, which displays up-regulated expression of Nrf2 target genes in an uninduced state. In this paper, we determine that the gene responsible for it768 was the zebrafish homolog of phosphomannomutase 2 (Pmm2), which is a key enzyme in the initial steps of N-glycosylation, and its mutation in humans leads to PMM2-CDG (congenital disorders of glycosylation), the most frequent type of CDG. The pmm2it768 larvae exhibited mild defects in N-glycosylation, indicating that the pmm2it768 mutation is a hypomorph, as in human PMM2-CDG patients. A gene expression analysis showed that pmm2it768 larvae display up-regulation of endoplasmic reticulum (ER) stress, suggesting that the activation of Nrf2 was induced by the ER stress. Indeed, the treatment with the ER stress-inducing compounds up-regulated the gstp1 expression in an Nrf2-dependent manner. Furthermore, the up-regulation of gstp1 by the pmm2 inactivation was diminished by knocking down or out double-stranded RNA-activated protein kinase (PKR)-like ER kinase (PERK), one of the main ER stress sensors, suggesting that Nrf2 was activated in response to the ER stress via the PERK pathway. ER stress-induced activation of Nrf2 was reported previously, but the results have been controversial. Our present study clearly demonstrated that ER stress can indeed activate Nrf2 and this regulation is evolutionarily conserved among vertebrates. Moreover, ER stress induced in pmm2it768 mutants was ameliorated by the treatment of the Nrf2-activator sulforaphane, indicating that Nrf2 plays significant roles in the reduction of ER stress.
The Keap1–Nrf2 system senses various environmental stresses and induces cytoprotective genes to protect cells from these stresses (13). The main components of the system are Nrf2 and Keap1; Nrf2 heterodimerizes with small Maf proteins and functions as a transcription activator while Keap1 represses Nrf2 functions as an adaptor for Cul3-based E3 ligase to regulate the proteasomal degradation of Nrf2. In addition to the Nrf2-inhibiting effects, Keap1 also functions as a sensor molecule for a variety of Nrf2-activating compounds. One of the most interesting points in the Keap1–Nrf2 system is that it can respond to cellular stresses, such as autophagy impairment and mitochondria stress (4, 5), in addition to well-studied oxidative stress and electrophiles.
The Keap1–Nrf2 system is well conserved among vertebrates, including zebrafish (68), in light of its molecular basis (911), activating chemicals and stress (1215), target genes (1618), and physiological functions (15, 19). We previously isolated six mutant zebrafish lines in which Nrf2 is activated without treatment with Nrf2 activators (12). We concluded that the mutations in these lines generate some endogenous cellular stresses that can activate Nrf2. The analysis of these mutants will help clarify the novel Nrf2-inducing cellular stresses and elucidate the molecular basis of their regulation.
In this study, we identified the gene responsible for it768, one of the six mutant lines, and elucidated the molecular mechanism for how the mutation activates the Keap1–Nrf2 system. The gene responsible for it768 was phosphomannomutase 2 (pmm2), which encodes the enzyme required for the biosynthesis of mannose-1-phosphate, an essential metabolite for an early step in the N-glycosylation process. pmm2it768 homozygous larvae showed impaired and insufficient N-glycosylation, followed by an increase in the endoplasmic reticulum (ER) stress. We found that the ER stress induces the up-regulation of the Keap1–Nrf2 system through the double-stranded RNA-activated protein kinase (PKR)-like ER kinase (PERK) pathway. The Nrf2 activation by ER stress has been demonstrated previously using cultured cells (20), but whether or not this is part of general regulation is controversial, since negative data were also reported by other groups (2123).

Results

pmm2 Is the Gene Responsible for it768.

it768 mutant embryos and larvae showed no obvious defects during embryogenesis and hatching process in bright-field images while the gstp1 expression in the gills was up-regulated in 5-d-postfertilization (dpf) larvae (Fig. 1A). The up-regulation of other Nrf2 targets, such as hmox1a, was also observed in the liver of 7-dpf larvae so Nrf2 may have been activated constitutively in the mutants. The expression of gstp1 and hmox1a in the mutant larvae was reduced when nrf2a was knocked down by morpholino oligonucleotide injection (nrf2aMO) (Fig. 1A), suggesting that up-regulation of these genes is indeed mediated by the Keap1–Nrf2 system. To confirm the Nrf2 dependence of the up-regulation, we crossed the it768 mutant line with the Nrf2 mutant line nrf2afh318 (19) and observed the gstp1 expression in pmm2it768;nrf2afh318 compound larvae. As expected, the nrf2a mutation decreased the it768-dependent gstp1 up-regulation (Fig. S1), which reflected the result of the nrf2a-knockdown analysis.
Fig. 1.
Up-regulated expression of Nrf2 target genes in it768 larvae that have splicing defects in the pmm2 gene. (A) Expression of gstp1 and hmox1a in WT or pmm2it768 larvae injected or not with 1 pmol nrf2aMO. The closed and open arrowheads indicate the gills and liver, respectively. The numbers in each picture indicate the larvae exhibiting strong expression of gstp1 or hmox1a/tested larvae. (B) Nucleotide and deduced amino acid sequences of the major cDNA isolated from WT and pmm2it768 larvae. The red “a” indicates the G-to-A mutation site of the it768 mutant in the zebrafish pmm2 genome. The green “R” indicates the amino acid residue corresponding to Arg141 in human PMM2. EcoRI indicates the EcoRI site only present in the WT cDNA.
To identify the gene responsible for it768, we restricted its chromosomal location to an ∼0.1-centimorgan interval on chromosome 1 using about 2,000 meioses and found pmm2 as a candidate (Fig. S2A). Phosphomannomutase is a highly conserved enzyme that catalyzes the conversion of mannose-6-phospate to mannose-1-phosphate, and, in vertebrates, two homologs—Pmm1 and Pmm2—are present (24). The amino acid sequence of the deduced zebrafish Pmm2 protein shared 70.4% identity with human PMM2 protein (Fig. S2B). In the zebrafish genome, one Pmm1 and one Pmm2 gene exist (www.ensembl.org/Danio_rerio/Info/Index). To confirm whether the up-regulation of gstp1 expression was based on the pmm2 mutation, we next performed a gene knockdown analysis using morpholino oligonucleotides (Fig. S2C). As expected, gstp1 was up-regulated in the WT AB larvae injected with an antisense morpholino oligonucleotide against pmm2 (pmm2MO), but not in those injected with the 5-mismatched type (5mis-pmm2MO). Conversely, the overexpression of pmm2 reduced the up-regulated expression of gstp1 in pmm2it768 homozygous larvae (Fig. S2D). These results indicated that the mutation of pmm2 is indeed the reason why the gstp1 expression was up-regulated in it768 mutants.
Genomic sequencing of pmm2 demonstrated that there were no marked differences in the coding region between WT and it768 mutants, but the G in the beginning of intron 5 was A in the mutant (Fig. 1B). Since it is the first G in the GU–AG rule (25), a splicing defect was suggested. We therefore next analyzed the pmm2 mRNA in WT and mutant larvae by reverse transcription-PCR (RT-PCR) and found that the WT mRNA was a bit longer than that from the it768 larvae, which lost the corresponding sequences to exon 5, including the EcoRI site (Fig. S2E). To investigate this splicing defect in detail, we subcloned cDNA from 2-dpf embryos and 5-dpf larvae and determined their nucleotide sequences (Fig. S2F). Normal type was not observed in the homozygous mutants; instead, four types of abnormal pmm2 cDNAs with different splicing types were identified, including “+3,” which is probably translated into enzymatic-active proteins. We therefore considered that pmm2it768 is not the null but hypomorphic mutant. Noteworthy, most of the isolated cDNA were the 30-nucleotides-deleted type (Δ30) in the exon 5 (total 94%, n = 90). Since Δ30 was an in-frame deletion, the resulting it768-type Pmm2 protein was deduced to have an internal 10-aa deletion (Fig. 1B). Importantly, the amino acid residue corresponding to Arg141 in human PMM2 was included in the deleted 10 amino acids (Fig. 1B, green R). Arg141 is a crucial residue for substrate binding (26) and highly conserved among Pmm proteins, including invertebrate Pmm proteins (Fig. S2G). Its mutation to histidine (R141H) is the most frequent mutation in PMM2-CDG (congenital disorders of glycosylation) (previously CDG-Ia) patients (27). CDG is a subset of genetic disorders characterized by defective N-glycosylation of serum and cellular proteins (2830). Since the R141H mutation in the recombinant human PMM2 proteins was shown to disrupt its enzymatic activity (31, 32), this 10-aa deletion will also eliminate the activity of the zebrafish Pmm2 protein.

pmm2it768 Larvae Displays N-Glycosylation Defects.

Pmm2 homozygous knockout mice were reported to be embryonically lethal (33), and, in humans, the lack of patients homozygous for the R141H mutation was shown to be highly statistically significant (34, 35). To determine whether or not pmm2 is essential for zebrafish development, the survival of larvae generated from heterozygous cross-breeding was examined (Fig. 2A). The results indicated that pmm2it768 homozygous larvae become weak after 7 dpf and die within 2 wk after fertilization whereas sibling WT and heterozygous mutants remain viable.
Fig. 2.
Lethality and glycosylation defects in pmm2it768 larvae. (A) The survival of pmm2it768 mutants calculated using the Kaplan–Meier method (78). (B) Mannose-related metabolic pathways in N-glycan synthesis. On the ER membrane, 14 sugars [2 N-acetylglucosamine (GlcNAc), 9 mannose (Man), and 3 glucose (Glc)] from nucleotide-activated sugars (UDP-GlcNAc, GDP-Man, Dol-P-Man, and Dol-P-Glc) are sequentially transferred to the lipid-like precursor dolichol phosphate (Dol-P) to synthesize the lipid-linked oligosaccharides (LLO) required for the N-glycosylation of proteins. PMM2 is an essential enzyme for generating mannose-1-phosphate (Man-1-P), which is required for the formation of both GDP-Man and Dol-P-Man. Tunicamycin (TM) blocks the transfer of GlcNAc to Dol-P. (C) Pyridylamino-glycans derived from the zebrafish larvae. The glycan content in WT and pmm2it768 was calculated based on the peak areas of the chromatograms on the amide column. (D) The HPLC profiles on the amide column of N-linked glycans from whole bodies of WT (black) or pmm2it768/it768 larvae (pink) at 5 dpf. MX (X = numbers) indicates ManxGlcNAc2-asn, and a, b, and c indicate pauci-mannose–type glycans. (E) N-glycosylation profiles on the octadecylsilyl column of fraction c separated on the amide column. The N-glycan structures were identified using the HPLC mapping method, as described in Materials and Methods. circles, mannose; squares, N-acetylglucosamine. Of note, the peak c1 was not detected in WT larvae.
Pmm2 is an essential enzyme for synthesizing oligosaccharides required for N-glycosylation (Fig. 2B) (28). To examine whether or not N-glycosylation was decreased in pmm2it768 larvae, the N-glycosylated protein extracted from 5-dpf larvae was analyzed. The total amount of N-linked oligosaccharides was reduced in pmm2it768 homozygous larvae compared with WT; instead, free oligosaccharide levels were increased (Fig. 2C and Fig. S3A). N-glycans were processed and modified in the secretory pathway (28). To investigate which steps in this processing were affected by the pmm2 mutation, we analyzed the N-glycosylated chains using HPLC and found that all high-mannose-type glycans were down-regulated (Fig. 2 C and D), suggesting that defects do not influence the processing steps in the Golgi body but much earlier steps, probably at the ER. In addition, we determined the structures of pauci-mannose–type glycans in peaks a, b, and c in Fig. 2D (Fig. 2E and Fig. S3 B and C) and found that abnormal-type N-glycans were accumulated in pmm2it768 larvae (Fig. 2E). Given these findings, we hypothesized that an extraordinary structure of N-glycans was generated in pmm2it768 larvae because of the shortage of GDP-mannose, resulting in the underglycosylation and accumulation of aberrant processed N-glycans.

Up-Regulation of Physiological ER Stress in the Liver of pmm2it768 Larvae.

The expression profile of pmm2 mRNA during the embryonic and larval stages was ubiquitous (Fig. S4A). Intriguingly, the pmm2 expression in the gills and liver became strong in pmm2it768 homozygous mutants compared with WT siblings after 3 dpf (Fig. S4B). Since the expression of the Pmm2 gene has been reported to be induced by ER stress in both mice and zebrafish (36, 37), we hypothesized that the up-regulation of pmm2 expression in pmm2it768 larvae was due to the induction of ER stress by the pmm2 mutation through the unfolded protein response (UPR) pathway. The UPR is a cellular protective response to ER stress, such as the transcriptional induction of the chaperone gene BiP/Grp78/Hspa5 and transcription factor Chop/Ddit3 (38, 39). Given that ER stress is induced when abnormal N-glycosylation occurs (40), we speculated that ER stress was up-regulated in pmm2it768 larvae. We therefore examined the expression of the ER stress/UPR marker bip/grp78/hspa5 (37, 41) in the mutant larvae by whole-mount in situ hybridization (WISH) and RT-PCR and found that the bip expression was up-regulated in the liver in pmm2it768 homozygous larvae (Fig. 3 AC). Another ER stress/UPR marker, chop/ddit3 (42, 43), was also up-regulated in the homozygous mutants (Fig. 3 B and C). These results indicate that ER stress was induced in the cells in the mutant liver.
Fig. 3.
PERK-dependent activation of Nrf2 in 5-dpf larvae by genetic- and chemical-induced ER stress. The numbers in each picture indicate the larvae exhibiting strong bip expression in the liver (open arrowheads)/tested larvae (A) or strong gstp1 expression in the gills (closed arrowheads)/tested larvae (DG). (A) bip expression in WT sibling and pmm2it768 larvae. (B) The expression of bip and chop was analyzed by RT-PCR in WT, pmm2it768/+ (m/+), or pmm2it768/it768 (m/m) larvae. The amount of cDNA used for RT-PCR was standardized by the ef1α expression. (C) The relative expression levels of bip and chop to ef1α was evaluated by qPCR in WT, pmm2it768/+ (m/+), or pmm2it768/it768 (m/m) larvae. a and b indicate statistically significant differences (Tukey’s test, P < 0.001; n = 3 for each genotype). (D) Induced expression of gstp1 after 12-h treatment of 5 μg/mL tunicamycin (TM) or 2 μM thapsigargin (TG) in WT AB larvae injected or not with 1 pmol nrf2aMO. (E) Induced expression of gstp1 after 12-h treatment of 1 μM thapsigargin (TG) in WT sibling and nrf2afh318 larvae. (F) gstp1 expression in WT sibling and pmm2it768 larvae injected or not with 1 pmol perkMO. (G) gstp1 expression in WT sibling and perkit312 larvae injected or not with 1 pmol pmm2MO. The graph shows the percentages of gstp1-positive larvae of the indicated genotypes. The numbers of larvae tested are indicated above the bars. Asterisks denote statistical significance (Fisher’s exact test; *P < 0.001). The data were obtained based on the results of the WISH analysis shown in the Upper panels.
Nrf2 has been shown to be activated by chemically induced ER stress in some mammalian cultured cells (44). The genetic ER stress induced by the pmm2 mutation may also activate zebrafish Nrf2. To test this hypothesis, we examined whether the Nrf2 target gene gstp1 is induced by treatment with the chemical ER stress inducers tunicamycin (TM) and thapsigargin (TG) in WT AB larvae (Fig. 3D). Tunicamycin is a competitive inhibitor of N-acetylglucosamine transferase that blocks protein N-glycosylation, similar to the pmm2 mutation (Fig. 2B), while thapsigargin is an inhibitor of the Ca2+-ATPase in the ER that induces perturbation of ER calcium homeostasis (45). As expected, gstp1 induction was observed in both tunicamycin- and thapsigargin-treated larvae, indicating that Nrf2 is activated by ER stress regardless of its underlying mechanisms, although the induction was weak and required a longer time than that induced by the typical Nrf2 activator diethyl maleate (DEM) (Fig. S5A). The induction was demonstrated to be Nrf2-dependent since it was eliminated when nrf2a was knocked down by morpholino oligonucleotides (Fig. 3D). The Nrf2 dependence was further confirmed by experiments using the Nrf2 mutant line nrf2afh318 (Fig. 3E). In contrast, the tunicamycin-induced bip expression was not reduced by nrf2a knockdown, suggesting that it was Nrf2-independent (Fig. S5B). This idea is also supported by the result that no bip induction was observed by diethyl maleate treatment (Fig. S5B).
Cullinan and Diehl (44) reported that the ER stress-induced activation of Nrf2 was mediated by the PERK pathway, out of three major UPR pathways (46). We therefore performed a knockdown analysis of perk/eif2ak3, the only ortholog of mammalian PERK in zebrafish (Fig. S6A), using splicing inhibition-type morpholino oligonucleotides. With nonspliced perk mRNA, the expression of the perk-dependent ER-stress marker chop (47) was drastically reduced while that of the perk-independent ER-stress marker bip was enhanced (Fig. S6B). This latter observation may suggest that the ER stress was up-regulated by perk knockdown. When perkMO-injected larvae were treated with tunicamycin or thapsigargin, the induction of gstp1 expression was obviously weakened, compared with the control larvae (Fig. S6C). No effects of perk knockdown on the diethyl maleate-induced expression of gstp1 were noted (Fig. S6C). Similarly, the up-regulated expression of gstp1 in pmm2it768 homozygous larvae was reduced by perk knockdown (Fig. 3F), suggesting that the ER stress-induced activation of Nrf2 was mediated by the PERK pathway. To strengthen our conclusion, we generated a perk-knockout zebrafish line (perkit312) by the CRISPR-Cas9 system (Fig. S6D) and performed gene expression analysis of gstp1 using pmm2-knockdown perkit312 larvae. As expected, the up-regulated expression of gstp1 in pmm2 morphants was suppressed by perk knockout (Fig. 3G). Taken together, these results indicate that Nrf2 was activated by ER stress in zebrafish larvae, at least in part, in a PERK-dependent manner.

Attenuation of ER Stress by the Nrf2 Activation.

The fact that the Keap1–Nrf2 system is activated in response to ER stress implies that this cellular defense system can suppress ER stress and/or its downstream adverse effects. To test this hypothesis, the expression of the ER stress marker bip in pmm2it768 larvae was analyzed in pmm2it768;nrf2afh318 compound larvae. Interestingly, the expression of bip in pmm2it768/it768;nrf2afh318/fh318 larvae was higher than in pmm2it768/it768;nrf2afh318/+ and pmm2it768/it768;nrf2a+/+ larvae and was elevated in accordance with the decrease in the WT nrf2a allele, suggesting that Nrf2 attenuated the ER stress in pmm2it768/it768 single mutants (Fig. S7A). We also examined the lethality of pmm2it768;nrf2afh318 compound larvae (Fig. S7B). The nrf2a mutation had no effect on the survival of pmm2it768 homozygous larvae, suggesting that Nrf2 is not directly related to either the lethality or survival of pmm2it768 larvae.
The finding of Nrf2-dependent suppression of ER stress suggested that further activation of the Keap1–Nrf2 system by exogenous chemical inducers might reduce the up-regulated ER stress in pmm2it768 larvae. To examine this possibility, pmm2it768 larvae were treated with sulforaphane, a well-known Nrf2 activator, and the expression of bip was analyzed. Fig. 4 indicates that sulforaphane treatment reduced the numbers of bip-positive larvae in pmm2it768 homozygous mutants, suggesting that Nrf2 activation can attenuate the pmm2 mutation-induced up-regulation of ER stress. As expected, the sulforaphane-dependent attenuation of bip up-regulation in the pmm2it768 homozygous larvae was canceled by the Nrf2 mutation (Fig. 4), suggesting that the effect of sulforaphane depends on the activation of Nrf2. Taken together, these results suggest that the Keap1–Nrf2 system plays a critical role in the UPR to reduce ER stress.
Fig. 4.
The Nrf2-dependent attenuation of the up-regulated ER stress by sulforaphane treatment. The graphs show the percentages of bip-positive 5-dpf larvae of the indicated genotypes and their expression after 12-h treatment of 40 μM sulforaphane (SF). The numbers of larvae tested are indicated above the bars. An asterisk denotes statistical significance (Fisher’s exact test; *P < 0.001). The data were obtained based on the results of the WISH analysis shown in the Right panels. The strength of the bip staining was determined by double-blind scoring. The arrowheads indicate the liver.

Discussion

In this study, we demonstrated that Nrf2 is activated by the genetic or chemical-induced ER stress via the PERK pathway and plays a critical role in reducing ER stress in zebrafish (Fig. S8). Some previous reports showed that ER stress activates Nrf2 (20, 48, 49) while others denied this regulation (2123). In addition to the latter reports, there were no transcriptome papers, which clearly showed the induction of Nrf2 target genes by ER stress. The ER stress-induced activation of Nrf2 has therefore not been widely recognized. In the present study, we clearly showed that ER stress activated Nrf2 in animals by genetic analysis and suggest that this regulation has been evolutionarily conserved among vertebrates. Previous studies may have failed to observe this regulation for one of two reasons: (i) The Nrf2 activation by ER stress is weak and slow compared with that by electrophiles or oxidative stress, as shown in Fig. S5A (TM, TG vs. DEM); and (ii) this regulation may depend on certain conditions, such as tissue specificities and developmental stages. Nrf1, a paralog protein of Nrf2, has been shown to exist in the ER and undergo N-glycosylation (5052). It is possible that Nrf1 was activated in response to ER stress and cooperated with Nrf2 to induce the expression of Nrf2 target genes, but this was not the case. The expression of gstp1 in both WT and pmm2it768 larvae was up-regulated, not down-regulated, by double knockdown of nrf1a and nrf1b (53), zebrafish coorthologs of mammalian Nrf1, suggesting that the gstp1 up-regulation in pmm2it768 larvae may not be mediated by Nrf1 (Fig. S9).
Attenuation of ER stress by Nrf2 activation suggests the importance of the Keap1–Nrf2 system in the homeostasis of cells producing a number of secreted factors. Indeed, it was reported that the tunicamycin-induced ER stress was worsened in an insulinoma cell line by Nrf2 knockdown (49). In the present study, the treatment of sulforaphane, a cancer chemopreventive ingredient in broccoli sprouts (5456), reduced the up-regulated ER stress in mutant larvae. Since the conservation of ER stress regulation among vertebrates has been validated using medaka fish and zebrafish (37, 57, 58), the attenuation of ER stress by Nrf2 activation may be an effective strategy, not only for human health, but also farming animals. The molecular basis of this attenuation is unknown, but enhancement of ER-associated degradation (ERAD) of unfolded and misfolded proteins by Nrf2-dependent up-regulation of proteasome subunits might have contributed (18, 49, 59).
Human PMM2 is known as the gene responsible for PMM2-CDG (60). In PMM2-CDG, the most common type of CDG, patients have a life-threatening liver insufficiency, with an overall 20% mortality during the neonatal period, combined with psychomotor retardation, hypotonia, dysmorphic features, failure to thrive, coagulopathy, abnormal endocrine functions, and a pronounced susceptibility to infection (61). No therapeutic options are available for PMM2-CDG patients at present (34). A trial to develop a zebrafish model of PMM2-CDG has recently been started using pmm2-knockdown zebrafish (62), but the phenotypes of their morphants were not observed in our pmm2it768 mutants: dysmorphic craniofacial cartilage and motility defects in the embryonic period (Table S3). The cause of this discrepancy is unclear. Since pmm2it768 is a hypomorphic mutant with splicing defects, it may display weaker symptoms than knockdown embryos. It is also possible that the phenotypic difference between pmm2it768 mutants and pmm2 morphants is due to off-target effects of morphant phenotypes (63) and/or genetic compensation by the deleterious mutation (64). As shown in Fig. 2, pmm2it768 larvae showed defects in N-glycosylation, as in PMM2-CDG patients (65, 66), suggesting that pmm2it768 larvae can be used as a disease model for PMM2-CDG. We have therefore attempted to search for chemical compounds that can suppress the lethality of pmm2it768 larvae, hoping to find drug candidates for PMM2-CDG, but no compound has been shown to be effective so far, including sulforaphane. We think that the cause of larval lethality may be hypoglycosylation of critical proteins for survival, not the up-regulated ER stress. Furthermore, the up-regulation of mild ER stress, which was detected in pmm2it768, has also been observed in some CDG patients (67). In contrast, the UPR was not detected in fibroblasts derived from CDG patients (68), probably because those fibroblasts produce fewer secreted proteins than liver cells, which expressed bip in our study. Since the cerebellum, one of the major organs in which CDG symptoms are observed, has been shown to be sensitive to ER stress (69), ER stress may be the cause of CDG symptoms in some patients. For such patients, diet therapy using Nrf2-activating foods/drinks/supplements, such as broccoli sprouts, may improve their conditions.

Materials and Methods

Zebrafish and Chemicals.

AB and TL strains were used as WT zebrafish. For genotyping pmm2it768, the genomic DNA at the mutation sites was amplified by PCR using primers in Table S1 and digested with ClaI: pmm2it768 mutant alleles were digested; WT alleles were not. Genotyping of nrf2afh318 was carried out as described previously (19). For the induction experiments and survival assays, larvae were placed in culture dishes with E3 medium plus 0.1 mg/L methylene blue (19), containing diethyl maleate (Wako), sulforaphane (LKT Laboratories), tunicamycin (Sigma-Aldrich), and/or thapsigargin (Sigma-Aldrich). In the case of survival assays, the E3 medium was exchanged every 2 d. Dead embryos were sequentially collected each day and genotyped as described above.

Chromosome Mapping.

The gene locus of the it768 mutant was roughly mapped by a bulked-segregant analysis using 333 polymorphic Z-markers (70). Fine mapping with additional polymorphic markers was carried out using ∼2,000 homozygous mutant larvae in the AB/TL hybrid F1 generation.

Plasmid Construction.

Full-length cDNA of pmm2 was synthesized by RT-PCR using zebrafish larval RNA and specific primers (Table S1) and subcloned into the pBluescript II KS and pCS2FL vectors (10); the resulting plasmids were named pKSpmm2 and pCS2FLpmm2, respectively. To construct pKSbip, the bip cDNA was synthesized by RT-PCR using specific primers (Table S1) and subcloned into the pBluescript II KS. The plasmids pSKhmox1a (17) and pKSgstp1N (16) have been described previously.

Gene Expression Analysis.

The WISH and RT-PCR analyses were carried out as described previously (14). To prepare RNA probes for the WISH analysis, pKSpmm2, pKSbip, and pKSgstp1N were digested with BamHI, and pSKhmox1a was digested with XhoI, mixed with DIG RNA labeling mix (Roche), and transcribed with T3 RNA polymerase (Roche). All pictures were taken using a Leica MZ16 microscope equipped with an Olympus DP73 digital camera. Genotyping of the stained embryos was carried out after taking photos. The specific primers used for the RT-PCR analysis are described in Table S1. A quantitative PCR (qPCR) was performed using a 7500 Fast Real-Time PCR System (Thermo Fisher Scientific) with THUNDERBIRD SYBR qPCR Mix (Toyobo). The expression level of each gene was normalized to the level in ef1α (or eef1a1l1). To visualize the splicing defects of pmm2, we amplified the region corresponding to exons 5 and 6 by RT-PCR using specific primers (Table S1) and digested the PCR products with EcoRI.

Microinjection.

Morpholino oligonucleotides (0.5 to 1 pmol) or capped mRNAs (100 pg) were injected into single cell-stage embryos using an IM300 microinjector (Narishige) as described previously (9). The nucleotide sequences of the morpholino oligonucleotides (Gene Tools) are listed in Table S2. The pmm2 mRNA was transcribed from pCS2FLpmm2 linearized by Bsp120I using the mMESSAGE mMACHINE kit (Ambion).

Preparation and Identification of Oligosaccharides and N-Glycans.

Ninety (WT) and 100 (pmm2it768) zebrafish larvae were lysed in hypotonic buffer, followed by homogenization and centrifugation to isolate free oligosaccharides, cytosolic proteins, and membrane-associated proteins as described previously (71). The N-glycans were released from the lyophilized protein fraction (soluble plus membrane-associated proteins) by hydrazinolysis and purified using a graphite carbon column (72). For the HPLC analysis, the reducing ends of the free and N-linked oligosaccharides were pyridylaminated and purified using a cellulose column (73, 74). The structures of N-glycans were identified by the HPLC mapping method combined with mass spectrometry. All of the experimental procedures used, including the chromatographic conditions and glycosidase treatments, have been described previously (75). The calculated HPLC map based on the unit contribution values was used for the estimation of a pauci-mannose–type pyridylamino-oligosaccharides (76).

Gene Knockout Using CRISPR-Cas9.

A knockout line of zebrafish perk was generated using CRISPR-Cas9 technology as previously described (77). In brief, transactivating CRISPR RNA (100 pg), perk-specific CRISPR RNA (50 pg), and Cas9 nuclease (400 pg) were coinjected into the yolk of one cell-stage WT AB embryos. All RNAs and protein were purchased from Integrated DNA Technologies. Identification of knockout line was performed by heteroduplex mobility assay and/or DNA sequencing using PCR and primers shown in Table S1.

Acknowledgments

We thank M. Eguchi, M. Komeda, Y. Nakayama, Y. Terashita, and Y. Wada for help with fish maintenance; T. E. Dever, H. H. Freeze, T. Irimura, T. Ishikawa, A. Kawahara, A. Kobayashi, K. Mori, H. Yoshida, and Y. Wada for valuable suggestions; and C.-S. Andrea, H. Nakajima, Miki Takeuchi, Miho Takeuchi, and J. Tamaoki for experimental help and discussions. This work was supported by grants from the Yamazaki Spice Promotion Foundation (to M.K.), the Koyanagi Foundation (to M.K.), the Japan Science and Technology Corporation (ERATO) (to M.Y.), and the Ministry of Education, Science, Sports and Culture of Japan [Grants 20059004, 21026003, 21659043, 24590340, 26116705, and 26520101 (to M.K.); Grant 25102008 (to K.K.); and Grants 15K07935 and 26110716 (to H.Y.)].

Supporting Information

Supporting Information (PDF)

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Information & Authors

Information

Published in

Go to Proceedings of the National Academy of Sciences
Proceedings of the National Academy of Sciences
Vol. 115 | No. 11
March 13, 2018
PubMed: 29472449

Classifications

Submission history

Published online: February 22, 2018
Published in issue: March 13, 2018

Keywords

  1. ER stress
  2. Nrf2
  3. PMM2-CDG
  4. sulforaphane
  5. zebrafish mutant

Acknowledgments

We thank M. Eguchi, M. Komeda, Y. Nakayama, Y. Terashita, and Y. Wada for help with fish maintenance; T. E. Dever, H. H. Freeze, T. Irimura, T. Ishikawa, A. Kawahara, A. Kobayashi, K. Mori, H. Yoshida, and Y. Wada for valuable suggestions; and C.-S. Andrea, H. Nakajima, Miki Takeuchi, Miho Takeuchi, and J. Tamaoki for experimental help and discussions. This work was supported by grants from the Yamazaki Spice Promotion Foundation (to M.K.), the Koyanagi Foundation (to M.K.), the Japan Science and Technology Corporation (ERATO) (to M.Y.), and the Ministry of Education, Science, Sports and Culture of Japan [Grants 20059004, 21026003, 21659043, 24590340, 26116705, and 26520101 (to M.K.); Grant 25102008 (to K.K.); and Grants 15K07935 and 26110716 (to H.Y.)].

Notes

This article is a PNAS Direct Submission.

Authors

Affiliations

Department of Molecular and Developmental Biology, Faculty of Medicine, University of Tsukuba, 305-8575 Tsukuba, Japan;
Present address: Department of Neuroanatomy and Embryology, School of Medicine, Fukushima Medical University, 960-1295 Fukushima, Japan.
Tadayuki Tsujita1
Department of Molecular and Developmental Biology, Faculty of Medicine, University of Tsukuba, 305-8575 Tsukuba, Japan;
Exploratory Research for Advanced Technology Environmental Response Project, Japan Science and Technology Agency, University of Tsukuba, 305-8575 Tsukuba, Japan;
Present address: Department of Applied Biochemistry and Food Science, Saga University, 840-8502 Saga, Japan.
Department of Molecular and Developmental Biology, Faculty of Medicine, University of Tsukuba, 305-8575 Tsukuba, Japan;
Li Li1
Department of Molecular and Developmental Biology, Faculty of Medicine, University of Tsukuba, 305-8575 Tsukuba, Japan;
Present address: School of Life Science and Technology, Harbin Institute of Technology, 150080 Harbin, China.
Hirokazu Yagi
Graduate School of Pharmaceutical Sciences, Nagoya City University, Mizuho-ku, 467-8603 Nagoya, Japan;
Yuji Fuse
Department of Molecular and Developmental Biology, Faculty of Medicine, University of Tsukuba, 305-8575 Tsukuba, Japan;
Yaeko Nakajima-Takagi
Department of Molecular and Developmental Biology, Faculty of Medicine, University of Tsukuba, 305-8575 Tsukuba, Japan;
Present address: Department of Cellular and Molecular Medicine, Graduate School of Medicine, Chiba University, 260-8670 Chiba, Japan.
Koichi Kato
Graduate School of Pharmaceutical Sciences, Nagoya City University, Mizuho-ku, 467-8603 Nagoya, Japan;
Okazaki Institute for Integrative Bioscience, National Institutes of Natural Sciences, Okazaki, 444-8787 Aichi, Japan;
Institute for Molecular Science, National Institutes of Natural Sciences, Okazaki, 444-8787 Aichi, Japan;
Masayuki Yamamoto
Exploratory Research for Advanced Technology Environmental Response Project, Japan Science and Technology Agency, University of Tsukuba, 305-8575 Tsukuba, Japan;
Department of Medical Biochemistry, Tohoku University Graduate School of Medicine, Aoba-ku, 980-8575 Sendai, Japan
Department of Molecular and Developmental Biology, Faculty of Medicine, University of Tsukuba, 305-8575 Tsukuba, Japan;
Exploratory Research for Advanced Technology Environmental Response Project, Japan Science and Technology Agency, University of Tsukuba, 305-8575 Tsukuba, Japan;

Notes

6
To whom correspondence should be addressed. Email: [email protected].
Author contributions: K.M., T.T., V.T.N., L.L., M.Y., and M.K. designed research; K.M., T.T., V.T.N., L.L., H.Y., Y.F., Y.N.-T., K.K., and M.K. performed research; K.M., T.T., V.T.N., and L.L. analyzed data; and M.K. wrote the paper.
1
K.M., T.T., V.T.N., and L.L. contributed equally to this work.

Competing Interests

The authors declare no conflict of interest.

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    Nrf2 activation attenuates genetic endoplasmic reticulum stress induced by a mutation in the phosphomannomutase 2 gene in zebrafish
    Proceedings of the National Academy of Sciences
    • Vol. 115
    • No. 11
    • pp. 2539-E2665

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