Next Article in Journal
Molecular Interplay Between Non-Coding RNAs and Connexins and Its Possible Role in Cancer
Previous Article in Journal
Jasonia glutinosa (L.) DC.: Back in Our Pantries? A Review of Its Pharmacological Activity and Mechanisms of Action
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
IJMS
Volume 26
Issue 6
10.3390/ijms26062537
ijms-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Elf1 Deficiency Impairs Macrophage Development in Zebrafish Model Organism

by
Qianli Tan
1,
Jing Wang
2,
Yimei Hao
2,
Shizeng Yang
2,
Biao Cao
2,
Weijun Pan
1,2 and
Mengye Cao
2,*
1
Shanghai Jiao Tong University School of Medicine, Shanghai 200025, China
2
Shanghai Institute of Nutrition and Health, University of Chinese Academy of Sciences, Chinese Academy of Sciences, Shanghai 200031, China
*
Author to whom correspondence should be addressed.
Int. J. Mol. Sci. 2025, 26(6), 2537; https://doi.org/10.3390/ijms26062537
Submission received: 5 February 2025 / Revised: 28 February 2025 / Accepted: 10 March 2025 / Published: 12 March 2025
(This article belongs to the Section Molecular Biology)
Browse Figures
Versions Notes

Abstract

:
The Ets (E-twenty-six) family of transcription factors plays a critical role in hematopoiesis and myeloid differentiation. However, the specific functions of many family members in these processes remain largely underexplored and poorly understood. Here, we identify Elf1 (E74-like factor 1), an Ets family member, as a critical regulator of macrophage development in the zebrafish model organism, with minimal impact on neutrophil differentiation. Through morpholino knockdown screening and CRISPR/Cas9-mediated gene editing, we demonstrate that Elf1 is critical for macrophage development and tissue injury responses. Specific overexpression of dominant-negative Elf1 (DN-Elf1) in macrophages demonstrated a cell-autonomous effect on macrophage infiltration. Furthermore, the overexpression of cxcr4b, a gene downstream of Elf1 regulation and essential for cell migration and injury response, significantly rescued this defect, indicating Elf1 as a key regulator of macrophage function. Our findings shed light on the roles of Elf1 in macrophage development and injury response and also highlight zebrafish as a powerful model for immunity research.
Keywords:
ELF1; ETS; zebrafish; macrophage

1. Introduction

Macrophages, essential components in the innate immune system, originate from hematopoietic progenitors derived from different hematopoietic origins [1]. Macrophages play crucial roles in immune responses, maintaining homeostasis, and facilitating tissue regeneration following injury. These functions include producing chemotactic factors to recruit inflammatory cells, clearing cellular debris, and secreting growth factors to promote wound healing, processes that rely heavily on macrophage infiltration at the injury site [2]. Despite the critical role of macrophages in immunity and tissue repair, the transcriptional mechanisms that govern their development and function remain poorly understood.
Multiple transcription factor families, including MAF (musculoaponeurotic fibrosarcoma) [3], IRF (interferon regulatory factor) [4], PPAR (peroxisome proliferator-activated receptor) [5], C/EBP (CCAAT/enhancer-binding protein) [6], and Ets (E-twenty-six) [7], regulate macrophage differentiation and maturation, as well as diverse functional responses, acting independently as well as through intricate interactions. For instance, IRF8 interacts with the transcription factor C/EBPα to influence the differentiation of myeloid progenitors into monocytes rather than neutrophils [8]. MAFB, a member of the MAF family, is critical for monocyte-to-macrophage differentiation and the polarization process [9]. Additionally, interactions between IRF8 and MAFB have been shown to regulate the cell cycle process and the differentiation of the human monocyte cell line THP-1 into macrophages [10].
The Ets family, characterized by a conserved DNA-binding ETS domain, is one of the largest and most widespread families of transcription factors across metazoans. To date, 28 Ets members have been identified in humans, 27 in mice, and 31 in zebrafish [11]. The Ets family plays a critical role in diverse cellular processes, including hematopoiesis, migration, and cell cycle regulation [12]. Based on a distance and parsimony analysis of the conserved ETS domains, the Ets family can be divided into 13 subgroups, including ETV (ETS variant), ELF (E74-like factor), and SPI (spleen focus forming virus proviral integration) [13]. Among these, the SPI subfamily member PU.1 is required for the development of multiple hematopoietic lineages and is involved in macrophage polarization and activation [14,15]. The ETV subfamily member ETV1 has been reported to regulate macrophage polarization in tumors [16]. These findings underscore the need for further research to explore the broader functions of other Ets family members, particularly in macrophages.
E74-like factor 1 (ELF1), a member of the Ets transcription factor family, was initially isolated from a human T-cell library using a probe targeting the ETS domain of ETS1 [17]. Elf1 has since been shown to bind to the regulatory regions of several genes critical for immune cell development and activation, including GM-CSF (granulocyte–macrophage colony-stimulating factor) [18], CD4 (cluster of differentiation 4) [19], and c-Fes (feline sarcoma oncogene) [20]. Elf1 is highly expressed throughout thymocyte development and in all stages of maturing thymocytes and T cells [21], with Elf1 knockout mice exhibiting a significant reduction in NKT cells both in the thymus and peripheral tissues [22]. Research has also demonstrated that the overexpression of the ETS DNA-binding domain of ELF1 can selectively disrupt its normal function without affecting the activity of other ETS family members, such as ETS1 or ETS2 [23]. Despite its abundant expression in macrophage cell lines [21], the precise functional role of ELF1 in macrophages remains poorly characterized, highlighting a gap in understanding its potential regulatory functions in immune cell regulation.
The zebrafish genome exhibits significant homology with the human genome, making it an invaluable model for studying macrophage development and function. Similar to mammals, zebrafish hematopoiesis occurs in distinct waves from different anatomical sites, giving rise to various myeloid lineages, each regulated by specific genetic programs [24]. Previous studies have identified 31 ETS domain genes in the zebrafish genome through systematic analysis and demonstrated the functions of erg, fli1, and etsrp in hematopoiesis, vasculogenesis, and angiogenesis [25]. However, the role of Elf1 in zebrafish hematopoiesis remains unclear. Moreover, the optical transparency of zebrafish larvae enables high-resolution, real-time imaging of macrophage behaviors and interactions during the developmental process. Additionally, the advancement of gene editing technologies has further underscored the advantages of zebrafish as a model for genetic manipulation.
Given the complex transcriptional mechanisms governing macrophage regulation, it is likely that additional, yet unidentified, regulators are involved. To investigate this, we employed ATAC-seq and RNA-seq in zebrafish macrophages to systematically identify novel transcription factors that influence macrophage development. Our results demonstrate that Elf1 is a crucial regulator of macrophage development and their response to injury. By using morpholino-mediated knockdown and CRISPR/Cas9-mediated gene editing approaches, we investigate the effects of Elf1 deficiency on macrophage function in zebrafish larvae. Our findings reveal that Elf1 regulates macrophage activity through the transcriptional control of key downstream genes.

2. Results

2.1. Enrichment of Ets Family Binding Motifs in Zebrafish Macrophage-Accessible Chromatin

To identify transcription factors critical for early macrophage development, we performed bulk ATAC sequencing once on zebrafish macrophages at 3 days post fertilization (dpf). After assessing data quality and analysis (See Section 4 and Figure S1A), we observed a significant enrichment of the GGAA motif, a known binding site for Ets family members, with 9 of the top 10 enriched motifs (by p-value) belonging to this family (Figure 1A). In the enriched results, we identified several Ets family transcription factors reported to control macrophage fate in zebrafish, including PU.1 [26] and Fli1 [27,28]. These findings not only support our results but also highlight the critical role of the Ets family in macrophage development and function. Given the diversity of Ets family members, we next assessed the expression of these factors in zebrafish macrophages through bulk RNA sequencing with three replicates (Figure 1B). After evaluating data quality and analysis (See Section 4 and Figure S1B,C), we found that pu.1 and fli1 were prominently expressed in macrophages, while erg, etv1, and ehf showed minimal expression. Notably, ets1, ets2, gabpa, and elf1 exhibited significant expression (Figure 1B). However, their specific roles in macrophage biology remain poorly understood, warranting further investigation.

2.2. Selective Regulation of Macrophage Development by Elf1

To investigate the regulatory roles of those Ets family members in macrophage development, we prioritized transcription factors based on their elevated expression profiles and motif enrichment patterns in zebrafish. Through morpholino (MO)-mediated knockdown, we aimed to uncover the specific functional contributions of these Ets factors to macrophage differentiation and activity during early development. Our results showed that only the Elf1 knockdown significantly reduced brain macrophage numbers, from an average of 26 to 18, while the knockdown of other Ets family members did not have a similar effect (Figure 2A,B). In addition, the Elf1 knockdown led to a decrease in macrophage numbers in the caudal hematopoietic tissue (CHT), whereas the knockdown of other Ets family members showed no impact (Figure 2C,D). These findings indicate that Elf1 specifically governs macrophage development during early zebrafish embryogenesis.
Additionally, the Elf1 knockdown did not affect somite length or overall embryonic development (Figure S2A,B), indicating that its role is specifically restricted to macrophage differentiation. Importantly, no significant alterations were observed in neutrophil numbers following Elf1 depletion (Figure S2C,D), underscoring Elf1′s selective involvement in macrophage development within the myeloid lineage, rather than affecting broader myeloid cell differentiation processes.

2.3. Macrophage Reduction in Elf1 Crispants

To further elucidate the impact of Elf1 on macrophages, we employed multiple gRNAs to target elf1, successfully replicating the elf1 knockout phenotype. Our results showed a significant reduction in macrophage numbers in both the brain and CHT regions of elf1 crispants, similar to the phenotype observed in elf1 morphants (Figure 3A–C,F). This further corroborates the essential role of Elf1 in macrophage development.
Given the multi-wave nature of zebrafish hematopoiesis during embryogenesis, we extended our analysis to macrophage development at 1 dpf and 2 dpf (Figure 3C). At 1 dpf, macrophage numbers were significantly reduced (Figure 3D), and at 2 dpf, a similar reduction was observed in the posterior blood island (PBI) region (Figure 3C,E). Collectively, these results highlight Elf1′s pivotal regulatory role in the development of macrophages throughout the transition from primitive to definitive hematopoiesis.

2.4. Impaired Macrophage Response to Injury in Elf1 Crispants

Macrophages play a crucial role in tissue repair and regeneration, playing a pivotal role in responding to injury signals. Transcription factors that regulate macrophage development also modulate their functional responses, such as activation during injury. Given Elf1′s critical role in macrophage development, we hypothesized that it might also impact macrophage activation in response to injury. To test this, we employed a tail amputation model in zebrafish larvae, which is a well-established assay for studying macrophage behavior during injury-induced inflammation [29]. Two hours post-amputation, we performed long-term imaging of the tail region to observe macrophage infiltration. The results showed that macrophage infiltration in elf1 crispants was reduced compared to control zebrafish larvae throughout the observation period (Figure 4A,B). Notably, at 10 h post-amputation, macrophage numbers in the wounded area were reduced by 56% in elf1 crispants compared to the control group, further highlighting the indispensable role of Elf1 in macrophage activation and response to tissue injury (Figure 4B). These results underscore Elf1’s crucial involvement not only in macrophage differentiation but also in regulating their functional responses during tissue repair.

2.5. Dominant-Negative Elf1 Reduces Macrophage Infiltration in a Cell-Autonomous Manner

To test whether Elf1′s effects on macrophage function are cell-autonomous, we overexpressed a dominant-negative form of Elf1 (DN-Elf1) specifically in macrophages. Using Tol2-mediated transgenesis in Tg(mpeg1:Gal4FF/UAS:eGFPcaax) zebrafish larvae, we selectively disrupted Elf1 function in macrophages under the control of the macrophage-specific mpeg1 promoter (Figure S3A,B) [30]. In the tail amputation assay, we observed a significant reduction in macrophage infiltration at the injury site in DN-Elf1-expressing zebrafish larvae. After 10 h post-amputation, fewer than half the number of macrophages were observed infiltrating the injury site compared to controls (Figure 5A,B). These results confirm that Elf1′s influence on macrophage activation is cell-autonomous.

2.6. Cxcr4b Mediates Elf1-Regulated Macrophage Infiltration

Given the cell-autonomous role of Elf1 in macrophage activation, we next explored the potential downstream targets of Elf1 that could mediate its effects on macrophage function. Notably, we found that cxcr4b, the zebrafish homolog of cxcr4, is as a key player in this process. Cxcr4b is highly expressed in macrophages (Figure 6A), and we identified an Elf1-binding motif within the open chromatin regions of the cxcr4b genetic locus. Cxcr4 is well established as a critical regulator of inflammatory responses, particularly in macrophage recruitment and tissue repair [31]. Based on these findings, we hypothesized that Elf1 might regulate macrophage function through Cxcr4b. Consistent with this, the qPCR analysis revealed a significant reduction in cxcr4b mRNA levels in macrophages from elf1 crispants compared to controls, suggesting that cxcr4b is a downstream target of Elf1 (Figure 6B). To further test this hypothesis, we examined macrophage infiltration following tail amputation and observed a significant reduction in macrophage infiltration at the injury site following tail transection in cxcr4b-deficient zebrafish larvae. Approximately one-third fewer macrophages infiltrated the wound area compared to controls (Figure 6C,D). These results collectively support the hypothesis that Elf1 regulates the macrophage response to injury through Cxcr4b.

2.7. Rescue of Macrophage Infiltration via Cxcr4b Overexpression

To further validate the role of Cxcr4b in Elf1-mediated macrophage function, we conducted a rescue experiment. Given the significant reduction in macrophage infiltration in elf1 crispants, we hypothesized that restoring Cxcr4b expression could rescue the impaired macrophage response. To test this, similar to the DN-Elf1 group, we specifically overexpressed Cxcr4b in macrophages in elf1 crispants using the macrophage-specific mpeg1 promoter (Figure S4A,B). Remarkably, compared to elf1 crispants, the Cxcr4b overexpression group exhibited a notable increase in macrophage infiltration at the injury site (Figure 7A). Statistical analysis revealed a one-third increase in macrophage infiltration in the Cxcr4b overexpression group 10 h after tail amputation (Figure 7B). These findings confirm that Elf1 regulates macrophage response to injury by modulating Cxcr4b expression.

3. Discussion

In this study, we utilized ATAC-seq to reveal a notable enrichment of Ets family binding motifs within the chromatin-accessible regions of zebrafish macrophages, underscoring the critical role of Ets family transcription factors in macrophage development. Through morpholino knockdown screening, we first demonstrated that Elf1 selectively regulates macrophage development during the early zebrafish developmental process, without impacting neutrophil differentiation. Further validation with CRISPR/Cas9-mediated elf1 mosaic mutants confirmed its essential role in both macrophage development and function. By overexpressing a dominant-negative form of Elf1, we demonstrated that Elf1′s influence on macrophage activation is cell-autonomous. Additionally, we found that Elf1 regulates macrophage responses to injury by modulating cxcr4b gene expression. Collectively, these findings highlight Elf1 as a critical transcription factor that specifically governs macrophage development and immune responses, underscoring its pivotal role in macrophage biology.
Both morpholino-based knockdown [32] and CRISPR/Cas9-mediated crispants [33] offer significant advantages for large-scale screening, time efficiency, cost-effectiveness, and simplicity. Despite each approach having inherent limitations, our core findings remained robust. By incorporating rigorous control comparisons and observing consistent results across both morpholino-based knockdown and CRISPR/Cas9-mediated crispant approaches, we further strengthened the reliability and interpretability of our findings, ensuring the validity of our results.
The Ets family transcription factors identified in our ATAC-seq enrichment analysis, including PU.1, Fli1, and Elf1, confirm the potential of the ATAC-seq approach in uncovering transcription factors involved in zebrafish macrophage development. In particular, Elf1, as a newly identified member, offers a novel research direction due to its association with macrophage development, further validating the effectiveness and reliability of ATAC-seq in studying zebrafish macrophage development.
The Ets family members are regulated by intricate networks of transcription factors and form functional complexes with specific partners to control downstream gene expression. Elucidating the regulatory mechanisms of ELF1 is essential for a deeper understanding of its pivotal role in gene regulation and cellular function. Previous studies have established a solid foundation for our findings. For example, PU.1 is essential for myeloid lineage development [34], and its high expression favors macrophage differentiation over neutrophil differentiation [35]. Consistent with this, the knockdown of PU.1 in the human monocyte cell line THP-1 results in a concurrent reduction in ELF1 expression [36], suggesting that ELF1 functions downstream of PU.1 in modulating macrophage development. Additionally, ELF1 has been shown to regulate the expression of key genes integral to macrophage biology. Notably, cooperative interactions between ELF1 and AP-1 play an important role in regulating inducible GM-CSF gene expression following human T-cell activation [18]. The human FES gene, which contains a strong myeloid-cell-specific promoter, is regulated by Sp1, PU.1, and ELF1 [19]. Similarly, the promoter and the macrophage-specific promoters of the mouse and human CSF1R genes are activated in RAW264 cells through the action of ETS2 and ELF1 [37]. These genes are essential for macrophage differentiation, activation, migration, survival, and proliferation, as well as immune responses [38,39,40]. Through its regulation of these genes, ELF1 deepens our insight into the molecular mechanisms through which its deficiency disrupts macrophage development and activation, revealing critical pathways involved in macrophage-specific functions and immune system dysregulation. ELF1 provides a clearer understanding of the molecular mechanisms underlying defects in macrophage development and activation. This evidence underscores ELF1 as a crucial regulator of macrophage-specific functions. However, a comprehensive exploration of the signaling networks and regulatory pathways is crucial to fully elucidate ELF1’s role in the regulation of myeloid lineage development.
During zebrafish embryogenesis, myeloid development is evolutionarily conserved across mammals. Zebrafish hematopoiesis consists of three distinct waves, each originating from different anatomical locations and governed by unique genetic programs [41,42]. The first wave, termed primitive hematopoiesis, generates myeloid cells and erythrocytes before 1 dpf [43]. Our results demonstrate a reduction in macrophages throughout the body at 1 dpf (Figure 3C,D), suggesting that Elf1 is involved in regulating primitive hematopoiesis. The second wave, intermediate hematopoiesis, occurs in the posterior blood island (PBI) from 24 to 48 hpf. This short wave produces committed erythromyeloid progenitors (EMPs), which differentiate into both erythroid and myeloid lineages [44]. Notably, we observed a reduction in macrophages within the PBI at 2 dpf (Figure 3C,E), indicating that Elf1 may also regulate intermediate hematopoiesis. Finally, definitive hematopoiesis is marked by the emergence of hematopoietic stem cells (HSCs), which then migrate to the caudal hematopoietic tissue (CHT) for proliferation and differentiation [45]. The reduction in macrophages in the CHT in both elf1 morphants and crispants (Figure 2 and Figure 3) further underscores Elf1’s involvement in definitive hematopoiesis. Collectively, these findings suggest that Elf1 plays a pivotal role in regulating all three waves of hematopoiesis (Figure 3). However, further studies are essential to confirm these observations and unravel the detailed mechanisms underlying Elf1’s regulatory functions.
Macrophages are crucial for immune defense, tissue repair, and inflammation regulation [1]. Beyond its well-established role in macrophage development and injury response, ELF1 likely governs a broader spectrum of macrophage functions. For instance, ELF1 modulates OAS1, a gene crucial for antiviral responses following IFN stimulation [46], highlighting its potential role in immune regulation beyond macrophage activation. Additionally, Elf1 regulates cxcr4b, a gene that not only mediates macrophage and microglia responses to oncogene activation in the brain [47] but also plays an essential role in initiating the granuloma-associated proangiogenic program in zebrafish [48]. Given that Cxcr4b is a marker for M2 macrophages [49], ELF1 may influence macrophage polarization and tissue repair processes, broadening its regulatory role in macrophage biology. Further exploration of the regulation of macrophage function by ELF1 will be valuable for understanding the underlying mechanisms and exploring potential therapeutic applications.
Overall, our study identified a novel function of Elf1 in regulating macrophage development, mediated through its genetic downstream factor, cxcr4b. These findings provide new insights into Elf1’s broader role in myeloid lineage differentiation and its impact on immune system regulation.

4. Materials and Methods

4.1. Zebrafish Husbandry and Transgenic Lines

Zebrafish used in this study were raised using the ESEN system (Esen Technology Development Co., Ltd., Beijing, China) with the following conditions: circulating water temperature at 28.5–29 °C, pH between 6.5 and 7.5, and a light-dark cycle of 14 h of light followed by 10 h of darkness. The density was approximately 5 adult zebrafish per liter of water. Adult fish were fed twice daily with freshly hatched brine shrimp nauplii, in accordance with the guidelines of the Institutional Animal Care and Use Committee [50]. Transgenic lines used in this study include Tg(mpeg1:eGFP) for labeling macrophages and Tg(mpx:GFP) for labeling neutrophil, as described previously [30,51]. Tg(mpeg1:Gal4FF/UAS:mCherrycaax) and Tg(mpeg1:Gal4FF/UAS:eGFPcaax) lines were generated using the Tol2-mediated Gal4-UAS system [52,53]. Embryos were incubated at 28.5 °C in egg water [0.6% (w/v) baysalt and 0.01% (w/v) methylene blue], which was replenished daily. To prevent melanin pigmentation, embryos were incubated in egg water containing 0.045%1-phenyl-2-thiourea (PTU, Sigma-Aldrich, St. Louis, MO, USA) starting from 1 dpf, with daily water changes. For experiments using embryos prior to 48 hpf, the chorion was manually removed using fine forceps under a microscope (SZX10, Olympus, Tokyo, Japan). All animal care and experimental procedures were approved by the Animal Research Advisory Committee of the Shanghai Institute of Nutrition and Health, Chinese Academy of Sciences.

4.2. Sorting of Zebrafish Macrophages

Zebrafish macrophages were isolated for sequencing and quantitative PCR (qPCR) analysis. Selected embryos were washed with pre-chilled PBS, and the yolk sac tissue was removed by adding Ringer’s solution and gently pipetting the embryos. After centrifugation, embryos were dissociated with 0.25% trypsin (Sigma-Aldrich) at 30 °C. Once complete dissociation into single cells was confirmed under a microscope, digestion was terminated with 10% FBS in PBS (Sigma-Aldrich) and 2mM CaCl2 solution. The embryos were then washed with 1% BSA in PBS and prepared for sorting. Macrophages were isolated using the MoFlo Astrios EQ (Beckman Coulter, Brea, CA, USA). Control group embryo samples were prepared simultaneously in parallel for comparison.

4.3. ATAC and RNA Sequencing

For bulk ATAC sequencing, following cell sorting, sample quality was assessed using a cell counter (Thermo Fisher Scientific, Seattle, WA, USA). Only samples with ≥90% cell viability and ≤5% adhesion rate was subjected to subsequent operations. Nuclei were isolated from fresh samples, and nucleus morphology was examined under the microscope. Qualified nuclei were then translocated using the Nextera Kit (Illumina, San Diego, CA, USA). After purification, the samples were amplified, quality checked, and sequenced on the Illumina HiSeq platform (Illumina). The quality of raw reads was examined using FastQC (version 0.11.9). Clean reads were obtained by removing adaptor sequences before mapping. The clean reads were then aligned to reference genome sequences using the BWA program (version 0.7.17), with mapping statistics (Figure S1A). Unique mapped reads were used for Peak Calling via MASC2 (version 2.1.1), applying a q-value threshold of <0.05. Motif analysis was conducted with HOMER software (version 4.11) and matched with the existing Motif data in public databases. Due to the stringent requirements for nuclear quality and quantity in ATAC-seq, as well as the need for subsequent validation through a series of complementary assays, including RNA-seq and in vivo experiments, we performed a round of bulk ATAC-seq on zebrafish macrophages to identify transcription factors as the initial screening.
For bulk RNA sequencing, RNA samples were directly reversed amplified using SMARTer Ultra Low Input RNA for the Illumina Kit (Takara, Kyoto, Japan), followed by sequencing on the Illumina HiSeq 2500 platform (Illumina). The quality of raw sequencing reads was assessed using FastQC (version 0.11.9). Raw data were filtered using Seqtk (version 1.3) to remove adapter sequences, trim low-quality bases at 3′ ends, discard reads shorter than 25 bases, and eliminate ribosomal RNA reads. Genome comparison was performed with Hisat2 (version 2.0.4). And the number of reads was standardized for gene expression. The bulk RNA-seq experiment was performed in triplicate, and reproducibility among the three replicates was confirmed using Spearman’s correlation coefficient analysis (Figure S1B,C).

4.4. Morpholino Preparation and Microinjection

Morpholino antisense oligonucleotides (MOs) were designed and purchased from Gene Tools, LLC (Philomath, OR, USA). The ets1 MO was previously reported in [54]. To design MOs, Ensemble accession numbers for target mRNA transcripts were identified based on differentially expressed transcripts and SNPs via PCR in zebrafish macrophages. The targets confirmed include elf1 (ENSDART00000122369.4) and ets2 (ENSDART00000161007.1), while the gabpa MO was designed to target all functional transcripts containing the ETS domain. Before ordering, as recommended by Gene Tools, LLC (Philomath, OR, USA), the specificity of the experimental MOs was verified in silico using the Ensemble BLAST/BLAT program (https://www.ensembl.org/, accessed on 25 January 2021). The MO sequences were input as nucleotide queries to identify any potential unintended targets in zebrafish cDNAs (transcripts/splice variants). Additionally, a standard control (Ctrl) MO was purchased as a negative control [55].
The lyophilized MOs were resuspended in nuclease-free water to make 3 mM stock solutions and aliquoted into 5 μL tubes. For experimental use, the following MOs were microinjected at the one-cell stage of zebrafish embryos: control MO (0.5 mM,1 nl), elf1 MO (0.5 mM,1 nl), ets1 MO (0.5 mM,1 nl), ets2 MO (0.5 mM, 1 nl), and gabpa MO (0.25 mM, 1 nl). Higher concentrations caused deformities in zebrafish. The MO sequences are listed in Table S1.

4.5. Crispant Generation

The gene sequences of zebrafish elf1 (ENSDART00000122369.4) and cxcr4b (ENSDARG00000041959) were obtained from Ensembl (www.ensembl.org, accessed on 20 November 2022). Targeted sequences were designed using CCTop-CRISPR/Cas9 online predictor (https://cctop.cos.uni-heidelberg.de:8043/, accessed on 21 November 2022) based on the composite score [56]. The gRNA sequences were input as nucleotide queries to identify any potential unintended targets in the zebrafish genomic sequence, and results were verified using Ensembl online tools. The modified gRNAs were ordered from Genescript, LLC (Nanjing, China). Compared with in vitro transcribed sgRNAs, the modified EasyEdit sgRNAs exhibited improved stability, reduced cytotoxicity, and higher editing efficiency [57].
Previous studies have demonstrated that utilizing multiple gRNAs in Cas9 ribonucleoprotein (RNP) complexes improves gene disruption efficiency and enables first-generation (F0) mosaic mutant zebrafish (crispants) to recapitulate the phenotypes of germline knockouts [58]. In this study, we employed multiple gRNAs to target genes and replicate the knockout phenotypes. The gRNAs and Cas9 protein (New England Biolabs, Ipswich, MA, USA) were co-microinjected into one-cell-stage embryos at a final concentration of 5 μM and a volume of 1 nl. To verify successful mutagenesis, genomic DNA was extracted from 10 randomly selected larvae per experimental group (n = 10/group), and PCR was performed using 2X PCR Master Mix (Beyotime, Shanghai, China). PCR products were sequenced to confirm the presence of the desired gene mutations. The gRNA sequences and genotyping details are listed in Table S1.

4.6. Plasmid Construction and Microinjection

The dominant-negative form of Elf1 (amino acids 200–297, NP_571234.2, NCBI) and the cxcr4b coding sequence (ENSDART00000061499.7, Ensembl) were amplified using PCR from cDNA derived from zebrafish larvae. For the pTol2: UAS-DN-Elf1-mCherry construct, the DN-Elf1 fragment was cloned into a modified Tol2-UAS-mCherry backbone. For the pTol2: UAS-cxcr4b-2A-mCherry construct, the cxcr4b coding sequences, followed by a 2A linker and mCherry in-frame, were cloned into a modified Tol2-UAS backbone. Restriction endonucleases and the KOD high-fidelity DNA polymerase were obtained from New England Biolabs. Vectors were ligated to linear segments using the Seamless Cloning Kit (Beyotime). Plasmids were purified using the phenol-chloroform method, and their concentration and purity were assessed using a Nanodrop spectrophotometer (Thermo Scientific). Embryos were injected with 20 pg of plasmid together with 15 pg of transposase mRNA. The transposase mRNA was synthesized in vitro using the SP6 mMessage mMachine Transcription Kit (Thermo Scientific). Primers used for plasmid construction and genotyping are listed in Table S1.

4.7. Total RNA Extraction and qPCR

Total RNA was extracted from sorted macrophages using the Trizol reagent and reverse-transcribed into cDNA using the HiScript IV 1st Strand cDNA Synthesis Kit (Vazyme, Nanjing, China). RNA concentrations were measured using a Nanodrop spectrophotometer (Thermo Scientific), and RNA quality was assessed through gel electrophoresis. Quantitative PCR was performed using SYBR Green Real-time PCR Master Mix (TOYOBO, Osaka, Japan) on a QuantStudio™6 Flex instrument (Thermo Scientific) with the comparative Ct (ΔΔCt) method. For qPCR, a 10 µL reaction mixture was prepared; it contained 15–25 ng of cDNA, 5 µL of SYBR Mix, and 0.3–0.4 µL of the forward and reverse primer mix (10 µM each), with the remainder filled with autoclaved Milli-Q® water (Sigma-Aldrich). Gene expression levels were normalized to the endogenous control ef1a in zebrafish. The primers used for qPCR have been previously reported [36]. Data were analyzed using the ΔΔCt method. Primers used for qPCR analysis are listed in Table S1.

4.8. Imaging and Count

Confocal imaging of live transgenic zebrafish larvae was performed using the Zeiss LSM 880 inverted confocal microscope (Carl Zeiss, Oberkochen, Germany) or the Olympus IXplore inverted confocal microscope equipped with the Yokogawa CSU-W1 spinning disk unit (Olympus). Embryos were anesthetized with 0.03% Tricaine (Sigma-Aldrich) and mounted in 1% low-melt agarose for imaging. Image processing was carried out using Imaris 9.8.0 (Bitplane, Abingdon, Oxon, UK) software. For brain imaging, a depth of approximately 150 μm was captured, and macrophages in the eye region were excluded from the quantification. In the caudal hematopoietic tissue (CHT) region, macrophages were counted within a 500 μm area extending from the cloaca. In the tail region, macrophage numbers were quantified within 150 μm from the incision site.

4.9. Statistical Analysis

Statistical analysis was performed using GraphPad Prism 8 software (San Diego, CA, USA). A one-way analysis of variance (ANOVA) with Dunnett’s test was applied to assess morpholino screening results. The unpaired two-tailed Student’s t-test was used to assess the statistical significance of differences between two groups. The two-way ANOVA with Sidak’s tests was used for analysis of the tail amputation model results. Data are presented as mean ± standard error of the mean (S.E.M.). p values < 0.05 were considered statistically significant, with * p < 0.05, ** p < 0.01, *** p < 0.001, and **** p < 0.0001, and ns indicated no significance (p ≥ 0.05).

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/ijms26062537/s1.

Author Contributions

Conceptualization, Q.T. and M.C.; methodology, Q.T. and S.Y.; formal analysis, Q.T. and Y.H.; investigation, Q.T., J.W. and S.Y.; resources, W.P.; data curation, B.C.; writing—original draft preparation, Q.T.; writing—review and editing, W.P. and M.C.; visualization, Q.T.; supervision, M.C.; project administration, M.C.; funding acquisition, W.P. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the National Key R&D Program of China (2023YFA1802000), the Key Research Program of Frontier Sciences, Chinese Academy of Sciences (ZDBS-LY-SM010), the CAS Project for Young Scientists in Basic Research (YSBR-077), and Shanghai Science and Technology Innovation Action Plan for Basic Research Program (21JC1406300) to W.P.

Institutional Review Board Statement

The animal study protocol was approved by the Animal Research Advisory Committee of Shanghai Institute of Nutrition and Health (SINH), Chinese Academy of Sciences (CAS) and conducted in accordance with Institutional Animal Care and Use Committee guidelines (SINH-2024-PWJ-1).

Informed Consent Statement

Not applicable.

Data Availability Statement

The data that support the findings of this study are accessible upon reasonable request from the corresponding author.

Acknowledgments

We thank Xiang Miao and Lin Qiu from Shanghai Institute of Nutrition and Health for technical supports. We also acknowledge the members of W.P. laboratory for zebrafish husbandry and technical support.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Varol, C.; Mildner, A.; Jung, S. Macrophages: Development and tissue specialization. Annu. Rev. Immunol. 2015, 33, 643–675. [Google Scholar] [CrossRef]
  2. Vannella, K.M.; Wynn, T.A. Mechanisms of Organ Injury and Repair by Macrophages. Annu. Rev. Physiol. 2017, 79, 593–617. [Google Scholar] [CrossRef]
  3. Zhang, C.; Guo, Z.M. Multiple functions of Maf in the regulation of cellular development and differentiation. Diabetes Metab. Res. Rev. 2015, 31, 773–778. [Google Scholar] [CrossRef]
  4. Taniguchi, T.; Ogasawara, K.; Takaoka, A.; Tanaka, N. IRF family of transcription factors as regulators of host defense. Annu. Rev. Immunol. 2001, 19, 623–655. [Google Scholar] [CrossRef]
  5. Toobian, D.; Ghosh, P.; Katkar, G.D. Parsing the Role of PPARs in Macrophage Processes. Front. Immunol. 2021, 12, 783780. [Google Scholar] [CrossRef]
  6. Yamanaka, R.; Lekstrom-Himes, J.; Barlow, C.; Wynshaw-Boris, A.; Xanthopoulos, K.G. CCAAT/enhancer binding proteins are critical components of the transcriptional regulation of hematopoiesis (Review). Int. J. Mol. Med. 1998, 1, 213–221. [Google Scholar] [CrossRef]
  7. Sharrocks, A.D. The ETS-domain transcription factor family. Nat. Rev. Mol. Cell Biol. 2001, 2, 827–837. [Google Scholar] [CrossRef]
  8. Kurotaki, D.; Yamamoto, M.; Nishiyama, A.; Uno, K.; Ban, T.; Ichino, M.; Sasaki, H.; Matsunaga, S.; Yoshinari, M.; Ryo, A.; et al. IRF8 inhibits C/EBPα activity to restrain mononuclear phagocyte progenitors from differentiating into neutrophils. Nat. Commun. 2014, 5, 4978. [Google Scholar] [CrossRef]
  9. Hamada, M.; Tsunakawa, Y.; Jeon, H.; Yadav, M.K.; Takahashi, S. Role of MafB in macrophages. Exp. Anim. 2020, 69, 1–10. [Google Scholar] [CrossRef]
  10. Gažová, I.; Lefevre, L.; Bush, S.J.; Clohisey, S.; Arner, E.; De Hoon, M.; Severin, J.; van Duin, L.; Andersson, R.; Lengeling, A.; et al. The Transcriptional Network That Controls Growth Arrest and Macrophage Differentiation in the Human Myeloid Leukemia Cell Line THP-1. Front. Cell Dev. Biol. 2020, 8, 498. [Google Scholar] [CrossRef]
  11. Findlay, V.J.; LaRue, A.C.; Turner, D.P.; Watson, P.M.; Watson, D.K. Understanding the role of ETS-mediated gene regulation in complex biological processes. Adv. Cancer Res. 2013, 119, 1–61. [Google Scholar]
  12. Yang, Y.; Han, X.; Sun, L.; Shao, F.; Yin, Y.; Zhang, W. ETS Transcription Factors in Immune Cells and Immune-Related Diseases. Int. J. Mol. Sci. 2024, 25, 10004. [Google Scholar] [CrossRef]
  13. Laudet, V.; Hänni, C.; Stéhelin, D.; Duterque-Coquillaud, M. Molecular phylogeny of the ETS gene family. Oncogene 1999, 18, 1351–1359. [Google Scholar] [CrossRef]
  14. Scott, E.W.; Simon, M.C.; Anastasi, J.; Singh, H. Requirement of transcription factor PU.1 in the development of multiple hematopoietic lineages. Science 1994, 265, 1573–1577. [Google Scholar] [CrossRef]
  15. Song, G.; Zhang, Z.; Chen, Y.; Hou, W.; Zhong, W.; Zhou, Y.; Zhang, A.; Xu, Y. PU.1 induces tumor-associated macrophages promoting glioma progression through BTK-mediated Akt/mTOR pathway activation. Am. J. Cancer Res. 2024, 14, 1139–1156. [Google Scholar] [CrossRef]
  16. Guo, X.; Li, Y.; Wan, B.; Lv, Y.; Wang, X.; Liu, G.; Wang, P. ETV1 inhibition depressed M2 polarization of tumor-associated macrophage and cell process in gastrointestinal stromal tumor via down-regulating PDE3A. J. Clin. Biochem. Nutr. 2023, 72, 139–146. [Google Scholar] [CrossRef]
  17. Thompson, C.C.; Brown, T.A.; McKnight, S.L. Convergence of Ets- and notch-related structural motifs in a heteromeric DNA binding complex. Science 1991, 253, 762–768. [Google Scholar] [CrossRef]
  18. Wang, C.Y.; Bassuk, A.G.; Boise, L.H.; Thompson, C.B.; Bravo, R.; Leiden, J.M. Activation of the granulocyte-macrophage colony-stimulating factor promoter in T cells requires cooperative binding of Elf-1 and AP-1 transcription factors. Mol. Cell Biol. 1994, 14, 1153–1159. [Google Scholar]
  19. Wurster, A.L.; Siu, G.; Leiden, J.M.; Hedrick, S.M. Elf-1 binds to a critical element in a second CD4 enhancer. Mol. Cell Biol. 1994, v14, 6452–6463. [Google Scholar]
  20. Heydemann, A.; Juang, G.; Hennessy, K.; Parmacek, M.S.; Simon, M.C. The myeloid-cell-specific c-fes promoter is regulated by Sp1, PU.1, and a novel transcription factor. Mol. Cell Biol. 1996, 16, 1676–1686. [Google Scholar] [CrossRef] [PubMed]
  21. Bassuk, A.G.; Barton, K.P.; Anandappa, R.T.; Lu, M.M.; Leiden, J.M. Expression pattern of the Ets-related transcription factor Elf-1. Mol. Med. 1998, 4, 392–401.s. [Google Scholar] [CrossRef] [PubMed]
  22. Choi, H.J.; Geng, Y.; Cho, H.; Li, S.; Giri, P.K.; Felio, K.; Wang, C.-R. Differential requirements for the Ets transcription factor Elf-1 in the development of NKT cells and NK cells. Blood 2011, 117, 1880–1887. [Google Scholar] [CrossRef] [PubMed]
  23. Huang, X.; Brown, C.; Ni, W.; Maynard, E.; Rigby, A.C.; Oettgen, P. Critical role for the Ets transcription factor ELF-1 in the development of tumor angiogenesis. Blood 2006, 107, 3153–3160. [Google Scholar] [CrossRef]
  24. Xu, J.; Du, L.; Wen, Z. Myelopoiesis during zebrafish early development. J. Genet. Genom. 2012, 39, 435–442. [Google Scholar] [CrossRef]
  25. Liu, F.; Patient, R. Genome-wide analysis of the zebrafish ETS family identifies three genes required for hemangioblast differentiation or angiogenesis. Circ. Res. 2008, 103, 1147–1154. [Google Scholar] [CrossRef] [PubMed]
  26. Jin, H.; Li, L.; Xu, J.; Zhen, F.; Zhu, L.; Liu, P.P.; Zhang, M.; Zhang, W.; Wen, Z. Runx1 regulates embryonic myeloid fate choice in zebrafish through a negative feedback loop inhibiting Pu.1 expression. Blood 2012, 119, 5239–5249. [Google Scholar] [CrossRef]
  27. Liu, F.; Walmsley, M.; Rodaway, A.; Patient, R. Fli1 acts at the top of the transcriptional network driving blood and endothelial development. Curr. Biol. 2008, 18, 1234–1240. [Google Scholar] [CrossRef]
  28. Zhang, A.; Lu, J.; Feng, S.; Yu, H.; Yu, T.; Zhao, S.; Chen, K.; Huang, Z.; Xu, J.; Qu, J.Y.; et al. Fli1 acts in parallel with Pu.1 to control macrophage and neutrophil fate in zebrafish. J. Genet. Genom. 2024, 51, 359–362. [Google Scholar] [CrossRef]
  29. Li, L.; Yan, B.; Shi, Y.Q.; Zhang, W.Q.; Wen, Z.L. Live imaging reveals differing roles of macrophages and neutrophils during zebrafish tail fin regeneration. J. Biol. Chem. 2012, 287, 25353–25360. [Google Scholar] [CrossRef]
  30. Ellett, F.; Pase, L.; Hayman, J.W.; Andrianopoulos, A.; Lieschke, G.J. mpeg1 promoter transgenes direct macrophage-lineage expression in zebrafish. Blood 2011, 117, e49–e56. [Google Scholar] [CrossRef]
  31. Werner, Y.; Mass, E.; Ashok Kumar, P.; Ulas, T.; Händler, K.; Horne, A.; Klee, K.; Lupp, A.; Schütz, D.; Saaber, F.; et al. Cxcr4 distinguishes HSC-derived monocytes from microglia and reveals monocyte immune responses to experimental stroke. Nat. Neurosci. 2020, 23, 351–362. [Google Scholar] [CrossRef] [PubMed]
  32. Moulton, J.D. Making a Morpholino Experiment Work: Controls, Favoring Specificity, Improving Efficacy, Storage, and Dose. Methods Mol. Biol. 2017, 1565, 17–29. [Google Scholar]
  33. Bolukbasi, M.F.; Gupta, A.; Wolfe, S.A. Creating and evaluating accurate CRISPR-Cas9 scalpels for genomic surgery. Nat. Methods 2016, 13, 41–50. [Google Scholar] [CrossRef] [PubMed]
  34. Rhodes, J.; Hagen, A.; Hsu, K.; Deng, M.; Liu, T.X.; Look, A.; Kanki, J.P. Interplay of pu.1 and gata1 determines myelo-erythroid progenitor cell fate in zebrafish. Dev. Cell 2005, 8, 97–108. [Google Scholar] [CrossRef] [PubMed]
  35. DeKoter, R.P.; Singh, H. Regulation of B lymphocyte and macrophage development by graded expression of PU.1. Science 2000, 288, 1439–1441. [Google Scholar] [CrossRef]
  36. Calero-Nieto, F.J.; Wood, A.D.; Wilson, N.K.; Kinston, S.; Landry, J.R.; Göttgens, B. Transcriptional regulation of Elf-1: Locus-wide analysis reveals four distinct promoters, a tissue-specific enhancer, control by PU.1 and the importance of Elf-1 downregulation for erythroid maturation. Nucleic Acids Res. 2010, 38, 6363–6374. [Google Scholar] [CrossRef]
  37. Ross, I.L.; Yue, X.; Ostrowski, M.C.; Hume, D.A. Interaction between PU.1 and another Ets family transcription factor promotes macrophage-specific Basal transcription initiation. J. Biol. Chem. 1998, 273, 6662–6669. [Google Scholar] [CrossRef]
  38. Ushach, I.; Zlotnik, A. Biological role of granulocyte macrophage colony-stimulating factor (GM-CSF) and macrophage colony-stimulating factor (M-CSF) on cells of the myeloid lineage. J. Leukoc. Biol. 2016, 100, 481–489. [Google Scholar] [CrossRef]
  39. Hackenmiller, R.; Kim, J.; Feldman, R.A.; Simon, M.C. Abnormal Stat activation, hematopoietic homeostasis, and innate immunity in c-fes-/- mice. Immunity 2000, 13, 397–407. [Google Scholar] [CrossRef]
  40. Hume, D.A.; Caruso, M.; Ferrari-Cestari, M.; Summers, K.M.; Pridans, C.; Irvine, K.M. Phenotypic impacts of CSF1R deficiencies in humans and model organisms. J. Leukoc. Biol. 2020, 107, 205–219. [Google Scholar] [CrossRef]
  41. Ellett, F.; Lieschke, G.J. Zebrafish as a model for vertebrate hematopoiesis. Curr. Opin. Pharmacol. 2010, 10, 563–570. [Google Scholar] [CrossRef] [PubMed]
  42. Jing, L.; Zon, L.I. Zebrafish as a model for normal and malignant hematopoiesis. Dis. Model. Mech. 2011, 4, 433–438. [Google Scholar] [CrossRef] [PubMed]
  43. Herbomel, P.; Thisse, B.; Thisse, C. Zebrafish early macrophages colonize cephalic mesenchyme and developing brain, retina, and epidermis through a M-CSF receptor-dependent invasive process. Dev. Biol. 2001, 238, 274–288. [Google Scholar] [CrossRef]
  44. Bertrand, J.Y.; Kim, A.D.; Violette, E.P.; Stachura, D.L.; Cisson, J.L.; Traver, D. Definitive hematopoiesis initiates through a committed erythromyeloid progenitor in the zebrafish embryo. Development 2007, 134, 4147–4156. [Google Scholar] [CrossRef]
  45. Murayama, E.; Kissa, K.; Zapata, A.; Mordelet, E.; Briolat, V.; Lin, H.-F.; Handin, R.I.; Herbomel, P. Tracing hematopoietic precursor migration to successive hematopoietic organs during zebrafish development. Immunity 2006, 25, 963–975. [Google Scholar] [CrossRef]
  46. Larsen, S.; Kawamoto, S.; Tanuma, S.; Uchiumi, F. The hematopoietic regulator, ELF-1, enhances the transcriptional response to Interferon-β of the OAS1 anti-viral gene. Sci. Rep. 2015, 5, 17497. [Google Scholar] [CrossRef]
  47. Chia, K.; Mazzolini, J.; Mione, M.; Sieger, D. Tumor initiating cells induce Cxcr4-mediated infiltration of pro-tumoral macrophages into the brain. eLife 2018, 7, e31918. [Google Scholar] [CrossRef] [PubMed]
  48. Torraca, V.; Tulotta, C.; Snaar-Jagalska, B.E.; Meijer, A.H. The chemokine receptor CXCR4 promotes granuloma formation by sustaining a mycobacteria-induced angiogenesis programme. Sci. Rep. 2017, 7, 45061. [Google Scholar] [CrossRef]
  49. Nguyen-Chi, M.; Laplace-Builhe, B.; Travnickova, J.; Luz-Crawford, P.; Tejedor, G.; Phan, Q.T.; Duroux-Richard, I.; Levraud, J.-P.; Kissa, K.; Lutfalla, G.; et al. Identification of polarized macrophage subsets in zebrafish. eLife 2015, 4, e07288. [Google Scholar] [CrossRef]
  50. Kimmel, C.B.; Ballard, W.W.; Kimmel, S.R.; Ullmann, B.; Schilling, T.F. Stages of embryonic development of the zebrafish. Dev. Dyn. 1995, 203, 253–310. [Google Scholar] [CrossRef]
  51. Jin, H.; Huang, Z.; Chi, Y.; Wu, M.; Zhou, R.; Zhao, L.; Xu, J.; Zhen, F.; Lan, Y.; Li, L.; et al. c-Myb acts in parallel and cooperatively with Cebp1 to regulate neutrophil maturation in zebrafish. Blood 2016, 128, 415–426. [Google Scholar] [CrossRef] [PubMed]
  52. Asakawa, K.; Kawakami, K. The Tol2-mediated Gal4-UAS method for gene and enhancer trapping in zebrafish. Methods 2009, 49, 275–281. [Google Scholar] [CrossRef]
  53. Davison, J.M.; Akitake, C.M.; Goll, M.G.; Rhee, J.M.; Gosse, N.; Baier, H.; Halpern, M.E.; Leach, S.D.; Parsons, M.J. Transactivation from Gal4-VP16 transgenic insertions for tissue-specific cell labeling and ablation in zebrafish. Dev. Biol. 2007, 304, 811–824. [Google Scholar] [CrossRef]
  54. Pham, V.N.; Lawson, N.D.; Mugford, J.W.; Dye, L.; Castranova, D.; Lo, B.; Weinstein, B.M. Combinatorial function of ETS transcription factors in the developing vasculature. Dev. Biol. 2007, 303, 772–783. [Google Scholar] [CrossRef] [PubMed]
  55. Pan, W.; Pham, V.N.; Stratman, A.N.; Castranova, D.; Kamei, M.; Kidd, K.R.; Lo, B.D.; Shaw, K.M.; Torres-Vazquez, J.; Mikelis, C.M.; et al. CDP-diacylglycerol synthetase-controlled phosphoinositide availability limits VEGFA signaling and vascular morphogenesis. Blood 2012, 120, 489–498. [Google Scholar] [CrossRef] [PubMed]
  56. Labuhn, M.; Adams, F.F.; Ng, M.; Knoess, S.; Schambach, A.; Charpentier, E.M.; Schwarzer, A.; Mateo, J.L.; Klusmann, J.H.; Heckl, D. Refined sgRNA efficacy prediction improves large- and small-scale CRISPR-Cas9 applications. Nucleic Acids Res. 2018, 46, 1375–1385. [Google Scholar] [CrossRef] [PubMed]
  57. Fei, J.; Zhao, D.; Pang, C.; Li, J.; Li, S.; Qiao, W.; Tan, J.; Bi, C.; Zhang, X. Mismatch prime editing gRNA increased efficiency and reduced indels. Nat. Commun. 2025, 16, 139. [Google Scholar] [CrossRef]
  58. Wu, R.S.; Lam, I.I.; Clay, H.; Duong, D.N.; Deo, R.C.; Coughlin, S.R. A Rapid Method for Directed Gene Knockout for Screening in G0 Zebrafish. Dev. Cell 2018, 46, 112–125.e4. [Google Scholar] [CrossRef]
Ijms 26 02537 g001
Figure 1. Transcription factor analysis of zebrafish macrophages via ATAC-seq and RNA-seq. (A) Motif enrichment analysis in zebrafish macrophages at 3 days post fertilization (dpf). The analysis reveals a strong enrichment of Ets family motifs, with 9 of the top 10 motifs belonging to the Ets family. (B) Expression levels of Ets family members in zebrafish macrophages, as determined via RNA-seq. Bar plots represent the mean ± standard error of the mean (S.E.M.).
Figure 1. Transcription factor analysis of zebrafish macrophages via ATAC-seq and RNA-seq. (A) Motif enrichment analysis in zebrafish macrophages at 3 days post fertilization (dpf). The analysis reveals a strong enrichment of Ets family motifs, with 9 of the top 10 motifs belonging to the Ets family. (B) Expression levels of Ets family members in zebrafish macrophages, as determined via RNA-seq. Bar plots represent the mean ± standard error of the mean (S.E.M.).
Ijms 26 02537 g001
Ijms 26 02537 g002
Figure 2. Morpholino knockdown-mediated screening of Ets family members in zebrafish macrophage development. (A) Representative confocal images showing macrophages in the head region of Tg(mpeg1:eGFP) zebrafish larvae at 3 dpf following morpholino-mediated knockdown of Ets family members. (B) Quantification of macrophage numbers in the head regions of the transgenic zebrafish shown in (A). Data were analyzed using one-way ANOVA followed by Dunnett’s test, with n ≥ 20, and shown as mean ± S.E.M. (C) Representative confocal images showing macrophages in caudal hematopoietic tissue (CHT) at 3 dpf following morpholino-mediated knockdown of Ets family members. (D) Quantification of macrophage numbers in the CHT of the transgenic zebrafish shown in (C). Data were analyzed using one-way ANOVA followed by Dunnett’s test, with n ≥ 15, and shown as mean ± S.E.M. Statistical significance is indicated as ns (p ≥ 0.05), **** p < 0.0001. Scale bar: 50 μm.
Figure 2. Morpholino knockdown-mediated screening of Ets family members in zebrafish macrophage development. (A) Representative confocal images showing macrophages in the head region of Tg(mpeg1:eGFP) zebrafish larvae at 3 dpf following morpholino-mediated knockdown of Ets family members. (B) Quantification of macrophage numbers in the head regions of the transgenic zebrafish shown in (A). Data were analyzed using one-way ANOVA followed by Dunnett’s test, with n ≥ 20, and shown as mean ± S.E.M. (C) Representative confocal images showing macrophages in caudal hematopoietic tissue (CHT) at 3 dpf following morpholino-mediated knockdown of Ets family members. (D) Quantification of macrophage numbers in the CHT of the transgenic zebrafish shown in (C). Data were analyzed using one-way ANOVA followed by Dunnett’s test, with n ≥ 15, and shown as mean ± S.E.M. Statistical significance is indicated as ns (p ≥ 0.05), **** p < 0.0001. Scale bar: 50 μm.
Ijms 26 02537 g002
Ijms 26 02537 g003
Figure 3. Reduction in macrophages in elf1 crispants. (A) Representative confocal images showing macrophages in the head region in transgenic zebrafish at 3 dpf following CRISPR-mediated mosaic mutants. Scale bar: 50 μm. (B) Quantification of macrophage numbers within the brain regions of the transgenic zebrafish shown in (A). Unpaired, two-tailed Student’s t-test, with n ≥ 20, and results are shown as mean ± S.E.M. (C) Representative confocal images of macrophages in whole zebrafish larvae at 1, 2, and 3 dpf. Scale bar: 200 μm; imaging depth approximately 200 μm. (D–F) Quantification of macrophage numbers within specific regions of the transgenic zebrafish shown in (C), including the posterior blood island (PBI) and caudal hematopoietic tissue (CHT). Unpaired, two-tailed Student’s t-test for n ≥ 12, 18, and 9, respectively; results are shown as mean ± S.E.M. Statistical significance indicated as ** p < 0.01 and **** p < 0.0001.
Figure 3. Reduction in macrophages in elf1 crispants. (A) Representative confocal images showing macrophages in the head region in transgenic zebrafish at 3 dpf following CRISPR-mediated mosaic mutants. Scale bar: 50 μm. (B) Quantification of macrophage numbers within the brain regions of the transgenic zebrafish shown in (A). Unpaired, two-tailed Student’s t-test, with n ≥ 20, and results are shown as mean ± S.E.M. (C) Representative confocal images of macrophages in whole zebrafish larvae at 1, 2, and 3 dpf. Scale bar: 200 μm; imaging depth approximately 200 μm. (D–F) Quantification of macrophage numbers within specific regions of the transgenic zebrafish shown in (C), including the posterior blood island (PBI) and caudal hematopoietic tissue (CHT). Unpaired, two-tailed Student’s t-test for n ≥ 12, 18, and 9, respectively; results are shown as mean ± S.E.M. Statistical significance indicated as ** p < 0.01 and **** p < 0.0001.
Ijms 26 02537 g003
Ijms 26 02537 g004
Figure 4. Macrophage behavior following tail amputation in elf1 crispants. (A) Time-lapse images showing macrophage infiltration in controls and elf1 crispants at 3 dpf. Imaging of the caudal fin region was initiated two hours post-amputation. The white dashed line marks the position of transection. Scale bar: 50 μm. (B) Quantification of macrophage numbers within the damage regions shown in (A). Data were analyzed using two-way ANOVA followed by Sidak’s tests, with n ≥ 7, and shown as mean ± S.E.M. Statistical significance indicated as * p < 0.05 and ** p < 0.01.
Figure 4. Macrophage behavior following tail amputation in elf1 crispants. (A) Time-lapse images showing macrophage infiltration in controls and elf1 crispants at 3 dpf. Imaging of the caudal fin region was initiated two hours post-amputation. The white dashed line marks the position of transection. Scale bar: 50 μm. (B) Quantification of macrophage numbers within the damage regions shown in (A). Data were analyzed using two-way ANOVA followed by Sidak’s tests, with n ≥ 7, and shown as mean ± S.E.M. Statistical significance indicated as * p < 0.05 and ** p < 0.01.
Ijms 26 02537 g004
Ijms 26 02537 g005
Figure 5. Overexpression of a dominant-negative form of Elf1 (DN-Elf1) in macrophages disrupts their response to injury. (A) Time-lapse imaging of macrophage infiltration at 3 dpf in control zebrafish and zebrafish expressing DN-Elf1 specifically in macrophages. The background transgenic line used was Tg(mpeg1:Gal4FF/UAS:eGFPcaax). Imaging of the caudal fin region was initiated two hours post-amputation. The white dashed line marks the position of transection. Scale bar: 50 μm. (B) Quantification of macrophage numbers within the damage shown in (B). Data were analyzed using two-way ANOVA followed by Sidak’s tests, with n ≥ 7, and shown as mean ± S.E.M. Statistical significance indicated as ** p < 0.01, *** p < 0.001, and **** p < 0.0001.
Figure 5. Overexpression of a dominant-negative form of Elf1 (DN-Elf1) in macrophages disrupts their response to injury. (A) Time-lapse imaging of macrophage infiltration at 3 dpf in control zebrafish and zebrafish expressing DN-Elf1 specifically in macrophages. The background transgenic line used was Tg(mpeg1:Gal4FF/UAS:eGFPcaax). Imaging of the caudal fin region was initiated two hours post-amputation. The white dashed line marks the position of transection. Scale bar: 50 μm. (B) Quantification of macrophage numbers within the damage shown in (B). Data were analyzed using two-way ANOVA followed by Sidak’s tests, with n ≥ 7, and shown as mean ± S.E.M. Statistical significance indicated as ** p < 0.01, *** p < 0.001, and **** p < 0.0001.
Ijms 26 02537 g005
Ijms 26 02537 g006
Figure 6. Macrophage behavior following tail amputation in cxcr4b crispants. (A) Expression levels of cxcr4b in zebrafish macrophages, as determined via RNA-seq. Bar plots represent the mean ± S.E.M. (B) Relative expression of cxcr4b in macrophages of elf1 crispants compared to the control group at 3 dpf. Unpaired, two-tailed Student’s t-test results for n = 4 are shown as mean ± S.E.M. Statistical significance indicated as **** p < 0.0001. (C) Time-lapse images showing macrophage infiltration in controls and cxcr4b crispants at 3 dpf. Imaging of the caudal fin region was initiated two hours post-amputation. The white dashed line marks the position of transection. Scale bar: 50 μm. (D) Quantification of macrophage numbers within the damage regions shown in (C). Data were analyzed using two-way ANOVA followed by Sidak’s tests, with n ≥ 10, and shown as mean ± S.E.M. Statistical significance indicated as * p < 0.05, ** p < 0.01, and *** p < 0.001.
Figure 6. Macrophage behavior following tail amputation in cxcr4b crispants. (A) Expression levels of cxcr4b in zebrafish macrophages, as determined via RNA-seq. Bar plots represent the mean ± S.E.M. (B) Relative expression of cxcr4b in macrophages of elf1 crispants compared to the control group at 3 dpf. Unpaired, two-tailed Student’s t-test results for n = 4 are shown as mean ± S.E.M. Statistical significance indicated as **** p < 0.0001. (C) Time-lapse images showing macrophage infiltration in controls and cxcr4b crispants at 3 dpf. Imaging of the caudal fin region was initiated two hours post-amputation. The white dashed line marks the position of transection. Scale bar: 50 μm. (D) Quantification of macrophage numbers within the damage regions shown in (C). Data were analyzed using two-way ANOVA followed by Sidak’s tests, with n ≥ 10, and shown as mean ± S.E.M. Statistical significance indicated as * p < 0.05, ** p < 0.01, and *** p < 0.001.
Ijms 26 02537 g006
Ijms 26 02537 g007
Figure 7. Ectopic expression of cxcr4b partially rescues macrophage function in elf1 crispants. (A) Time-lapse images of macrophage infiltration in controls and cxcr4b ectopic expression in elf1 crispant zebrafish larvae at 3 dpf. Imaging of the caudal fin region was initiated two hours post-amputation. The white dashed line marks the position of transection. Scale bar: 50 μm. (B) Quantification of macrophage numbers within the damage regions shown in (A). Data were analyzed using two-way ANOVA followed by Sidak’s tests, with n ≥ 18, and shown as mean ± S.E.M. Statistical significance indicated as ** p < 0.01, *** p < 0.001, and **** p < 0.0001.
Figure 7. Ectopic expression of cxcr4b partially rescues macrophage function in elf1 crispants. (A) Time-lapse images of macrophage infiltration in controls and cxcr4b ectopic expression in elf1 crispant zebrafish larvae at 3 dpf. Imaging of the caudal fin region was initiated two hours post-amputation. The white dashed line marks the position of transection. Scale bar: 50 μm. (B) Quantification of macrophage numbers within the damage regions shown in (A). Data were analyzed using two-way ANOVA followed by Sidak’s tests, with n ≥ 18, and shown as mean ± S.E.M. Statistical significance indicated as ** p < 0.01, *** p < 0.001, and **** p < 0.0001.
Ijms 26 02537 g007
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Tan, Q.; Wang, J.; Hao, Y.; Yang, S.; Cao, B.; Pan, W.; Cao, M. Elf1 Deficiency Impairs Macrophage Development in Zebrafish Model Organism. Int. J. Mol. Sci. 2025, 26, 2537. https://doi.org/10.3390/ijms26062537

AMA Style

Tan Q, Wang J, Hao Y, Yang S, Cao B, Pan W, Cao M. Elf1 Deficiency Impairs Macrophage Development in Zebrafish Model Organism. International Journal of Molecular Sciences. 2025; 26(6):2537. https://doi.org/10.3390/ijms26062537

Chicago/Turabian Style

Tan, Qianli, Jing Wang, Yimei Hao, Shizeng Yang, Biao Cao, Weijun Pan, and Mengye Cao. 2025. "Elf1 Deficiency Impairs Macrophage Development in Zebrafish Model Organism" International Journal of Molecular Sciences 26, no. 6: 2537. https://doi.org/10.3390/ijms26062537

APA Style

Tan, Q., Wang, J., Hao, Y., Yang, S., Cao, B., Pan, W., & Cao, M. (2025). Elf1 Deficiency Impairs Macrophage Development in Zebrafish Model Organism. International Journal of Molecular Sciences, 26(6), 2537. https://doi.org/10.3390/ijms26062537

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop