SBDS Gene Mutation Increases ROS Production and Causes DNA Damage as Well as Oxidation of Mitochondrial Membranes in the Murine Myeloid Cell Line 32Dcl3

Abstract

Shwachman–Diamond syndrome (SDS) is an autosomal recessive disease caused by mutation in the Shwachman–Bodian–Diamond syndrome (SBDS) gene. SDS has a variety of clinical features, including exocrine pancreatic insufficiency and hematological dysfunction. Neutropenia is the most common symptom in patients with SDS. SDS is also associated with an elevated risk of developing myelodysplastic syndromes and acute myeloid leukemia. The SBDS protein is involved in ribosome biogenesis, ribosomal RNA metabolism, stabilization of mitotic spindles and cellular stress responses, yet the function of SBDS in detail is still incompletely understood. Considering the diverse function of SBDS, the effect of SBDS seems to be different in different cells and tissues. In this study, we established myeloid cell line 32Dcl3 with a common pathogenic SBDS variant on both alleles in intron 2, 258 + 2T > C, and examined the cellular damage that resulted. We found that the protein synthesis was markedly decreased in the mutant cells. Furthermore, reactive oxygen species (ROS) production was increased, and oxidation of the mitochondrial membrane lipids and DNA damage were induced. These findings provide new insights into the cellular and molecular pathology caused by SBDS deficiency in myeloid cells.

INTRODUCTION

Shwachman–Diamond syndrome (SDS) is a rare autosomal recessive disorder with neutropenia and congenital bone-marrow failure as its cardinal symptoms.¹⁾ Hematologic manifestations other than neutropenia include anemia, increased fetal hemoglobin levels, thrombocytopenia, and aplastic anemia.^1,2) SDS also exhibits a variety of clinical features, including short stature, exocrine pancreatic insufficiency, and predisposition to leukemic transformation.^1,2) Approximately 20 to 25% of patients develop acute myelogenous leukemia (AML).²⁾ Most patients with SDS have been reportedly found to harbor mutations in the Shwachman–Bodian–Diamond syndrome (SBDS) gene.³⁾ Approximately 90% of the affected individuals exhibit SBDS variants. The most common mutations were 258 + 2T > C, followed by 183–184TA > CT. The most common combination of mutations was 183–184TA > CT/258 + 2T > C (50%), 258 + 2T > C/missense mutation (27.8%), 183–184TA > CT, 258 + 2T > C/258 + 2T > C (5.1%), and 258 + 2T > C/258 + 2T > C (4.4%).³⁾ The mutations result in a premature stop-codon (K62X) (184TA > CT) and a splice-defect (258 + 2T > C) that causes marked decline of the SBDS protein in the patient’s cells.

The SBDS gene is highly conserved and its orthologs are found in diverse species, ranging from archaea to vertebrate animals.³⁾ Therefore, SBDS is considered to play a fundamental role in cellular processes. To date, SBDS has been suggested to be involved in ribosome biogenesis and ribosomal RNA metabolism.^4,5) Weis et al. showed that SBDS interacts with elongation factor-like guanosine 5′-triphosphatase 1 to release the ribosome anti-association factor eIF6 from the late cytoplasmic pre-60S ribosomal subunit, allowing assembly of the 80S subunit.⁵⁾ In addition, SBDS might play a role in multiple biological processes, including mitotic spindle stabilization, cellular stress responses, cell proliferation and differentiation.^6–10) Considering the variety of clinical features in SDS, the contribution by and the regulation of the SBDS protein seems to be different among different cells and tissues. However, little is known about the precise function of SBDS and hence also about the pathogenesis of SDS.

Here, to investigate cellular dysfunction in myeloid cells, we established mutant cells (SDS cells) with a common pathogenic SBDS variant on both alleles in intron 2, i.e., 258 + 2T > C in murine myeloid cell line 32Dcl3. We then examined the characteristic properties of the SDS cells and analyzed several types of cellular damage caused by the SBDS gene mutation in the cells.

MATERIALS AND METHODS

Materials

32Dcl3 cells were obtained from the Riken Cell Bank (Tsukuba, Japan). The anti-SBDS antibody (sc-271350) was obtained from Santa Cruz Biotechnology (Dallas, TX, U.S.A.). The anti-DYKDDDDK (FLAG) tag antibody (018-22381), anti-α-tubulin antibody (017-25031), and anti-β-actin antibody (281-98721) were from FUJIFILM Wako Pure Chemical Corporation (Osaka, Japan). The anti-phospho-Histone H2A.X antibody (9718) was from Cell Signaling Technology (Danvers, MA, U.S.A.). The anti-puromycin (3RH11) antibody was from Cosmo Bio (Tokyo, Japan). 2′,7′-Dichlorodihydrofluorescein diacetate (H₂DCFDA), MitoPeDPP, MitoBright LT Deep Red, and mtSOX Deep Red, Cell Counting Kit-8 were from Dojindo (Kumamoto, Japan). MitoTEMPO and MitoQ were from Selleck Chemicals (Houston, TX, U.S.A.).

Cell Culture

Interleukin-3 (IL-3) dependent 32Dcl3 and SDS cells were cultured in Iscove’s modified Dulbecco’s medium (IMDM) supplemented with 10% fetal calf serum (FCS) and 10% WEHI3b conditioned medium, which contained murine IL-3. Cell proliferation was determined by Cell Counting Kit-8 assay.

Preparation of 32Dcl3 Cells with SBDS Gene Mutation Harboring Cells (SDS Cells)

The nuclease for the exchange of the SBDS gene were constructed as follows. Design of the guide RNAs was carried out using the CRISPR Design Tool (http://crispr.mit.edu) to minimize potential off-target effects. crRNAs were obtained from Integrated DNA Technologies (IDT; Coralville, IA, U.S.A.) for the testing of the Alt-R system. The guide RNA sequence for SBDS c.258 + 2T > C was GCAAGCAGGUAGGUCCUGCC. Alt-R crRNA and Alt-R tracrRNA were mixed in an equimolar ratio and heated at 95 °C for 5 min. Then, annealed Alt-R gRNA was allowed to form at room temperature for 10 min. Alt-R Cas9 nuclease (IDT) was complexed with gRNA to form an RNP complex in phosphate-buffered saline (PBS). RNP complex and the donor DNA template CTCATCAGTGCATTTGGGACAGACGACCAGACTGAAATCTGCAAGCAGGCACGTCCTGCCACGTGCAATGTAACAAATCTCACGATGGTAGGCAACATCT was electroporated into 32Dcl3 cells. After the transfection of the RNP complexes, clonal cell lines were isolated by dilution. The mutation of the SBDS gene was confirmed using the Big Dye V.3.1 Terminator Kit in an ABI Prism 3130 sequencer.

SUnSET Assay

A protein synthesis assay was performed using the SUnSET method.¹¹⁾ Cells were incubated with 1 µM puromycin for 30 min. Finally, puromycin labeled peptides in the same amount of cell extract (25 µg protein) were detected by immunoblotting.

Detection of Reactive Oxygen Species (ROS) Production

Cells were resuspended in PBS and treated with 100 nM H₂DCFDA for 10 min at 37 °C with or without 5 mM N-acetylcysteine (NAC). The fluorescence of the oxidized dichloro fluorescein (DCF) was analyzed with a fluorescein-activated cell sorting (FACS) flow cytometer.

Analysis of Oxidized Mitochondrial Membrane Lipids

MitoPeDPP is a cell-membrane-permeable perylene-based dye. It specifically localizes in mitochondria due to the triphenylphosphonium moiety of the dye and is used to monitor lipid peroxidation. Cells were washed with Hank’s buffer containing 8 mM PIPES (pH 7.3) and treated with 1 µM MitoPeDPP. The fluorescence of the oxidized MitoPeDPP by various peroxides was detected using a fluorescence microscope. The excitation and emission wavelengths of MitoPeDPP were 452 and 470 nm, respectively.

Detection of DNA Damage

Histone H2AX is a member of the H2A histone family that contains an evolutionarily conserved SQ motif at the COOH-terminus. Ser139 within this motif becomes rapidly phosphorylated by ATM and ATR kinases to yield a form known as γH2AX in response to double-strand DNA breaks associated with DNA damage.^12,13) Cells were attached on slide glass treated with 1 mg/mL poly-L-lysine (Sigma). The cells were fixed with 4% paraformaldehyde in PBS and permeabilized with 0.2% Triton X-100 in PBS. The antibody against γH2AX was used as the primary antibody. FITC conjugated anti-mouse immunoglobulin G (IgG) antibody was used as the secondary antibody. The fluorescence was analyzed under fluorescence microscopy (Olympus Life Science).

Immunoblotting

The cells were lysed in extraction buffer (20 mM Tris–HCl, 150 mM NaCl, 1 mM ethylenediaminetetraacetic acid (EDTA), 1 mM Na₂VO₃, 1% NP-40, 1 µM leupeptin, 1 µM pepstatin A, 1 mM phenylmethylsulfonyl fluoride, pH 7.5). Lysates were centrifuged at 12000 × g for 15 min at 4 °C to remove debris, and protein concentrations were determined with the Lowry method. The cell extract (20 µg protein) was resolved with 12.5% sodium dodecyl sulfate-polyacrylamide gels and transferred to polyvinylidene difluoride (PVDF) membranes. The membranes blocked in 5% nonfat milk in TBS-T (200 mM NaCl, 20 mM Tris–HCl, 0.05% Tween 20, pH 5.0). The membranes were then incubated with primary antibodies and horseradish peroxidase-conjugated anti-immunoglobulin secondary antibodies. The antigen-antibody complex was detected by enhanced chemiluminescence.

Statistical Analysis

The data are expressed as the mean ± standard deviation, and statistical significance was assessed with Welch Two sample t-test and Tukey test. A value of p < 0.05 was considered statistically significant.

RESULTS

Establishment of SDS Cells Using the Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/CRISPR-Associated Protein 9 (Cas 9) System

To clarify the role of SBDS in 32Dcl3 cells, we prepared SDS cells with a mutation in the mouse SBDS locus using the CRISPR/Cas9 system. The mutation c.258 + 2T > C was confirmed by DNA sequencing (Fig. 1A). Three clones (SDS#1–3) were established, and these clones showed similar reduced proliferation (Fig. 1B). We used mainly SDS#1 cells in the following experiments. First, we examined expression of the SBDS protein in SDS cells. SBDS with a molecular mass of 32 kDa was detected in wild-type cells, while SBDS was hardly detected in SDS cells (Fig. 1C). These results suggest that the mutation causes a deficiency of the SBDS protein by a splice-defect in intron 2 of the SBDS gene in SDS cells.

Fig. 1. Characteristic Properties of SDS Cells

(A) Nucleotide sequence of wild-type and mutant SBDS. Sbds mutations were observed in all of the SDS clones. (B) Growth curve of control 32Dcl3 (wild-type) and SDS cells. Mean cell proliferation (from 3 independent experiments) of control wild-type cells and SDS cells. (C) Expression of the SBDS protein in wild-type and SDS cells. Each protein extract was subjected to immunoblot analysis. The membrane was probed with an antibody against SBDS. (D) Translation monitoring using puromycin-labeled proteins in wild-type and SDS cells. Cells were labeled with puromycin, and whole cell lysate was subjected to immunoblot analysis. The membrane was probed with an antibody against puromycin. Quantification was performed using ImageJ software (National Institutes of Health). We have conducted the experiment using three clones once. Then, we obtained similar results in separate experiments using SDS#1 (data not shown).

It is well known that SBDS involves ribosome maturation, and the cells derived from SDS patients display a low amount of mature ribosomes.¹⁴⁾ We analyzed protein synthesis in SDS cells using the SUnSET assay based on puromycin incorporation.¹¹⁾ The de novo protein synthesis capacity of SDS cells decreased to approximately 30% of that of control cells (Fig. 1D). Taken together, these results demonstrate that SDS cells can be used as a disease model.

The Mutation of the SBDS Gene Increases the ROS Level in SDS Cells

Previous studies have shown that knockdown of the SBDS protein increases the ROS levels in various cell lines such as HeLa cells and the human leukemic K562 cells.^15,16) To examine whether the ROS level is increased in the SDS cells, the cells were incubated with H₂DCFDA and the fluorescence level of the oxidized fluorescence compound DCF was analyzed by flow cytometry. As shown in Fig. 2A, the fluorescence level in SDS cells was increased compared to that of wild-type cells. To confirm whether the fluorescence was derived from ROS, the SDS cells were incubated with NAC, a free radical scavenger. As a result, the fluorescence level in SDS cells was decreased and became equal to that in wild-type cells treated with NAC (Fig. 2B). Furthermore, to confirm that the deficiency of SBDS is responsible for the increase of the fluorescence level, we transfected the SBDS gene encoding a FLAG tag at the NH₂-terminus of SBDS in SDS cells and examined the expression of the SBDS protein as well as the fluorescence level. As shown in Figs. 2C and D, a considerable amount of FLAG-SBDS was expressed in SDS cells and the fluorescence level in the cells decreased to the level of wild-type cells. These results suggest that the increase of the ROS level in SDS cells was indeed caused by the deficiency of the SBDS protein.

Fig. 2. Increased ROS Production in SDS Cells

(A) Detection of the ROS level in wild-type and SDS cells. Cells were cultured for three days, and the level of ROS production was measured using oxidized DCFDA and flow cytometry. (B) ROS inhibition by NAC treatment. Wild-type and SDS cells were cultured for three days in the presence or absence of NAC for the last two days. The ROS production levels were measured as in Fig. 2A. (C) Overexpression of FLAG-SBDS in wild-type and SDS cells. wild-type and SDS cells were transfected with either pcDNA3.1/FLAG-SBDS or an empty plasmid. The stably transfected cells were selected by culturing them for a few months with medium containing 1 mg/mL G418. The overexpression of FLAG-SBDS was confirmed by immunoblot analysis with an anti-SBDS antibody and anti-FLAG antibody. (D) The reduced ROS in SDS cells was determined by the overexpression of FLAG-SBDS. The ROS levels in wild-type and SDS cells stably transfected with FLAG-SBDS were compared to those of control wild-type and SDS cells.

Oxidative Damage of the Mitochondrial Membrane and Nuclear DNA

It is known that an excess of ROS leads to damage of subcellular organelles.^17,18) Therefore, we assessed peroxidation of mitochondrial membrane lipids using the fluorescent imaging dye MitoPeDPP that accumulates selectively in mitochondrial membranes. As shown in Figs. 3A and B, the relative intensity of oxidized MitoPeDPP was significantly increased in SDS cells compared to wild-type cells. In addition, treatment with NAC in SDS cells reduced the fluorescence intensity equivalent to that of wild-type cells (Supplementary Fig. 1), suggesting that the oxidation of mitochondrial membrane lipids was induced by ROS in SDS cells. We further analyzed whether mitochondrial superoxide production was increased in SDS cells. The fluorescence intensity of mtSOX Deep Red was significantly increased in SDS cells, suggesting that O₂⁻ is produced in mitochondria (Supplementary Fig. 2). Such an increased ROS level would be expected to induce mitochondrial membrane damage.

Fig. 3. Mitochondrial Membrane Damage in SDS Cells

(A, B) Mitochondrial membrane lipid oxidation in SDS cells. Wild-type and SDS cells were stained by MitoPeDPP to detect oxidized mitochondrial membrane lipids, and the images were detected by fluorescence microscopy. The fluorescence intensity per cell was measured using ImageJ software.

Concerning DNA damage, histone H2AX, a member of the histone H2A family, is phosphorylated on Ser139 during DNA double-strand breaks.¹³⁾ The phosphorylated H2AX is defined as γH2AX. Therefore, anti-γH2AX antibody may be used to indicate the sites of DNA damage. As shown in Fig. 4A, the number of fluorescence foci in SDS cells was higher than that in control wild-type cells. When the number of foci was calculated in individual cells, the average number in SDS cells was 7.7 ± 1.9, which was 1.63-fold higher than in wild-type cells (Fig. 4B). In addition, treatment with NAC reduced the number of foci in the SDS cells (Fig. 4C), suggesting that the DNA damage was induced by the ROS that were induced by SBDS deficiency in SDS cells.

Fig. 4. DNA Damage in SDS Cells

(A, B) Increased DNA damage foci in SDS cells. Wild-type and SDS cells were cultured for three days and the γH2AX foci were detected by immunofluorescent staining. The number of foci per cell was compared. (C) DNA damage foci decreased by NAC treatment. Wild-type and SDS cells were cultured for three days in the presence or absence of NAC for the last two days. The number of γH2AX foci per cell was compared.

DISCUSSION

SDS is a multisystem disorder caused by mutation of the SBDS gene.³⁾ SDS exhibits multiple organ dysfunction with a wide range of clinical severity. It is characterized by pancreatic insufficiency and bone marrow failure syndrome and displays a tendency to leukemic transformation. The SBDS protein is involved in ribosome processing and maturation.^4,5) In addition, SBDS is implicated in having additional functional roles, including stabilizing the mitotic spindle and cellular stress, as well as cell proliferation and differentiation.^6–10) However, the exact function of SBDS remains poorly elucidated and the medical treatment of SDS is thus limited to the alleviation of symptoms. Understanding the pathogenesis of AML is one of critical issues related to SDS. Myeloid cells proliferate and differentiate to neutrophils, eosinophils and basophils in the presence of certain cytokines. Impaired function of SBDS might result in neutropenia and/or leukemia.

The murine myeloid cell line 32Dcl3 is one of the few cell lines that can terminally differentiate into neutrophils.¹⁹⁾ Differentiated wild-type cells exhibit a striking morphologic similarity to normal neutrophils. In this study, we established 32Dcl3 cells with a common pathogenic SBDS variant and examined changes in cellular function using 32Dcl3 cells as the control. SDS cells grew as well as wild-type cells, although displayed reduced saturation density compared with the control wild-type cells (Fig. 1B). This phenomenon is similar to our previous observation with SBDS knockdown in 32Dcl3 cells.²⁰⁾ In addition, the proliferation of SDS cells was recovered by the overexpression of TagGFP-SBDS (Supplementary Fig. 3). On the other hand, the protein synthesis in SDS cells was markedly decreased by approx. 70% compared to wild-type cells (Fig. 1D), suggesting that SBDS deficiency causes dysfunction of ribosomes associated with impaired ribosomal RNA (rRNA) maturation in SDS cells. A substantial decrease in protein synthesis might lead to impaired proliferation and maturation of myeloid cells.

Concerning the cellular dysfunction that occurs in SDS cells, we first found that the ROS level increased using the membrane permeable fluorescence dye H₂DCFDA, which produces fluorescence after oxidation. The claim that the increased fluorescence is caused by ROS and deficiency of the SBDS protein is supported by the following evidences. The fluorescence level became the same as in the control as the result of treatment the free radical scavenger NAC (Fig. 2B). The expression of considerable amount of FLAG-SBDS in SDS cells reduced the fluorescence level to that of the control (Figs. 2C, D). In addition, mitochondrial superoxide production seems to be induced in SDS cells (Supplementary Fig. 2). Although we found that ROS production is increased in SDS cells, we have no knowledge of which pathway is responsible for increased ROS production. Dysregulation of the anti-oxidation system might occur in SDS cells. As protein synthesis is markedly reduced in SDS cells, the amount of antioxidative enzymes such as superoxide dismutase (SOD), catalase and glutathione peroxidase, as well as synthesis of glutathione and plasmalogen containing polyunsaturated fatty acids, might be decreased. In fact, it is reported that the amount of SOD2 in mitochondria was decreased in a yeast model of SDS.²¹⁾

One of targets of ROS is the cell membrane, especially the phospholipids with polyunsaturated fatty acids.²²⁾ We analyzed the oxidation of mitochondrial membrane lipids using MitoPeDPP, which accumulates selectively in the mitochondrial membrane. The intensity of oxidized MitoPeDPP was increased significantly in SDS cells (Fig. 3A). The fluorescence of MitoPeDPP was superimposed to that of MitoBright LT, a reagent for mitochondria staining, suggesting that mitochondrial lipids are oxidized (Supplementary Fig. 4). The average of the fluorescence in individual SDS cell increased 1.2-fold compared to control wild-type cells (Fig. 3B). Mitochondria are important for the supply of energy through oxidative phosphorylation, and damaged mitochondria are a critical feature of impaired cellular function. In addition, although oxidized phospholipids are normally removed through the hydrolysis carried out by platelet activating-factor (PAF) acetylhydrolase II,²³⁾ the accumulation of oxidized phospholipids is known to be involved in many different diseases.²⁴⁾ It is evidently important to characterize how oxidized phospholipids affect cellular function of myeloid cells. It has been reported that mitochondrial function is impaired in SDS model cells, and the hypothesis put forward that lipid oxidation of mitochondrial membranes is responsible for this mitochondrial dysfunction.²⁵⁾

DNA damage can affect the transmission of genetic information, and it can be induced by a variety of endogenous attacks, including attacks by free radicals.²⁶⁾ The DNA double strand break is associated with loss of genetic information due to erroneous DNA repair. We examined DNA double strand breaks using an antibody against anti-γH2AX recognizing DNA damage site. The evidences for increased DNA damage in SDS cells are as follows.²⁷⁾ The number of fluorescent foci was found to be increased in SDS cell WEs (Fig. 4A) and the average number in SDS cells was 1.63-fold higher than control wild-type cells (Fig. 4B). The number of γH2AX foci in SDS cells was not reduced by the overexpression of FLAG-SBDS (data not shown), whereas cell proliferation of SDS cells expressing TagGFP-SBDS was recovered (Supplementary Fig. 3).

As NAC treatment reduced DNA damage, we examined whether NAC recovered cell proliferation and protein synthesis in SDS cells. However, cell proliferation and protein synthesis in SDS cells were not recovered (Supplementary Fig. 5). The results suggest that the decreased cell growth and protein synthesis are simply not caused by DNA damage and that they are independent phenomena. In addition, we examined whether mitochondrial ROS production causes DNA damage. As shown in Supplementary Fig. 6, mitochondrial-targeted antioxidant reagents such as MitoTEMPO, MitoQ and Ferrostatin-1 did not rescue DNA damage, suggesting that DNA damage may be independent against ROS produced in mitochondria. In addition, several papers suggested side effects of MitoQ and MitoTEMPO on ROS production.^28–31) Although we do not know the reason at this time, one possible reason is that antioxidation in the nucleus may not be recovered in the cells. On the other hand, treatment with NAC reduced the number of foci in SDS cells (Fig. 4C). In SDS cells, ROS production was significantly higher, indicating that DNA damage had occurred. These findings suggest that errors accumulate as a result of inadequate DNA damage repair.

Taken together, this study shows that deficiency of the SBDS protein induced oxidative stress and increased ROS production in SDS cells. The production of ROS and ensuing damage to the mitochondria were observed. It is possible that ROS-induced DNA fragmentation and the separately reported abnormalities that occur during the division phase may contribute to the development of conditions such as AML. As the precise mechanisms underlying the entry into neutropenia and AML are as yet unknown, SDS cells will provide us with a critically needed tool to obtain useful information for understanding the cellular and molecular pathology of SDS in future study. It is hope that just such understanding will also provide insights into treatments for this debilitating condition.

Acknowledgments

This work was supported in part by Hiroshima International University Special Research Grant Program.

Conflict of Interest

The authors declare no conflict of interest.

Supplementary Materials

This article contains supplementary materials.

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