Introduction

Understanding the regulatory mechanism of gonadal development is the basis for achieving large-scale aquaculture production. Catches of cephalopods have remained at a high level over the past 20 years, reaching 3.7 million tons in 2020 (FAO 2022). Many efforts have been made to artificially breed cephalopods worldwide (Bo et al. 2014; Ibarra-García et al. 2018; Jiang et al. 2020; Meza‐Buendía et al. 2021; Rodríguez et al. 2006). However, current catches and production still do not meet the demand, so there is an urgent need to develop cephalopod cultivation.

The long-armed octopus, Octopus minor (Sasaki, 1920), is widely distributed in the coastal waters of China, Korea, and Japan (Norman et al. 2014). As an important and promising marine mollusk in China, O. minor is tasty and rich in protein and multiple unsaturated fatty acids (Qian et al. 2010; Zheng et al. 2014). To date, there have been several reports on the artificial culture of O. minor (Bo et al. 2014, 2016; Liu 2013; Nan 2020; Qian et al. 2013, 2016; Song et al. 2018; Wu and Lv 1996; Zheng et al. 2009). Unlike many octopod species, such as O. vulgaris and O. mimus, which have a planktonic period in their life history characterized by small eggs and high fecundity (Mangold 1983; Warnke 1999), O. minor has relatively large eggs and low fecundity, laying 9–125 eggs per female (Qian et al. 2013). To master the reproductive strategies of cephalopods, several studies have been done on gametogenesis (de Lima et al. 2013; López-Peraza et al. 2013; Sieiro et al. 2014; Wang et al. 2015). The morphology and histogenesis of gonads in O. minor have been reported (Li 2010; Qian et al. 2016; Xu et al. 2008), but the stage classification is still incomplete. Besides, our current understanding of the molecular reproductive profile of O. minor remains limited. Therefore, knowing the reproductive mechanisms is of prime importance for the artificial breeding of O. minor.

Omics technology has developed rapidly in recent years, offering a powerful tool for clarifying the molecular mechanisms of physiological biochemistry. Transcriptome can directly display gene expression and has been applied to study potential reproductive mechanisms of octopods in the optic lobe, white body, and gonad (Juarez et al. 2019; Lopez-Galindo et al. 2018; Ventura-Lopez et al. 2022; Wang and Ragsdale 2018). Whole genomes of nine cephalopods have been published so far (Jiang et al. 2022), providing precise molecular information. As for O. minor, only its whole genome study and transcriptome analysis of ammonia stress have been conducted (Kim et al. 2018; Xu and Zheng 2020), and no other omics research has been reported.

Low fecundity and long embryonic period are the two main factors that restrict O. minor breeding (Qian et al. 2013; Zheng et al. 2014). Figuring out the basic biology of gonads and the molecular developmental mechanisms will lay the foundation for the genetic mechanism of reproduction. Hence, this study investigated reproductive biology, containing the gonadal morphological features and histological process, and transcriptome analysis of O. minor, aiming to provide a molecular basis for octopus breeding.

Material and Methods

Sample Collection and Morphological Measuring

O. minor is neither an endangered nor protected species. All work followed the principles stated in the EU Directive 2010/63/EU guidelines on cephalopod use and was approved by the Institutional Animal Care and Use Committee of Ocean University of China.

From September 2020 to June 2021, a total of 210 lively and healthy O. minor samples were collected monthly using bottom trawl in Swan Lake (37.3°N, 122.6°E), Shandong Province, China. In addition, eleven juveniles with invisible gonads and a male juvenile weighing 27 g were collected at the same site in June 2023. For all samples with observable gonads, total body weight (TW) and gonad weight (GW) were recorded to the nearest 0.001 g. The gonad somatic index (GSI) of O. minor was calculated according to Wang et al. (2015): GSI = (GW/TW) × 100. Mean GSI values were generated separately for males and females.

Histological Examination and RNA Extraction

All octopuses were anesthetized in seawater-prepared 4% ethanol for 20 min based on a prior study (Butler-Struben et al. 2018). The sex of O. minor samples was identified by examining the right third arm, with only males having a spoon-like hectocotylus. The ovaries and testes were subsequently removed and divided into two parts. One part was fixed in Bouin’s solution and preserved in 70% ethanol for histological examination, and the other part was frozen in liquid nitrogen and stored at −80 ℃ for RNA extraction.

For histological analysis, fixed gonads were dehydrated with gradient alcohol, cleared by xylene, and then embedded in paraffin. Sections of 5-μm thickness were obtained using a Leica RM 2016 rotary microtome (Shanghai, China) and then stained with hematoxylin-eosin. Finally, the prepared sections were examined and photographed with an Olympus BX 53 microscope (Olympus, Tokyo, Japan). Different cell types in ovaries and testes were identified and described based on previous research (ICES 2010; Wang et al. 2015).

After microscopic observation, it was noticed that the ovary of O. minor remained invisible to the naked eye, despite the oocytes having developed to the late primary oocyte (LPO) stage. Furthermore, the transition from vitellogenic oocytes (VO) to advanced vitellogenic oocytes (AVO) required about half a year. Additionally, catching female O. minor with fully developed eggs in the wild is quite difficult since they stay in burrows awaiting spawning. In contrast, testes have already matured when they are visible to the naked eye, so we can generally only observe the mature and spent stages. Twelve samples were chosen for RNA sequencing, comprising three replicates of the ovaries at stages of VO and AVO, and three replicates of testes at stages of mature and spent. Ovaries at VO and AVO were collected in January and April 2021, respectively, while testes at mature and spent stages were collected independently in January and May 2021. Total RNA was extracted using TRIzol reagent following the manufacturer’s protocol. RNA concentration was measured by NanoDrop 2000 and RNA quality was assessed using Agilent 5400.

cDNA Library Construction and Sequencing

Total RNA from gonads was used as input material for sample preparations. Sequencing libraries were then generated using NEBNext® UltraTM RNA Library Prep Kit for Illumina® (NEB, USA) according to the manufacturer’s recommendations.

Fragmentation was carried out using divalent cations under elevated temperature in NEBNext First Strand Synthesis Reaction Buffer (5X). For the first strand, cDNA was synthesized by random hexamer primer and M-MuLV ReverseTranscriptase (RNase H). For the second strand, cDNA synthesis was subsequently performed using DNA Polymerase I and RNase H. Remaining overhangs were converted into blunt ends by exonuclease/polymerase activities. Then 3′ ends of DNA fragments were adenylated, and NEBNext Adaptors with hairpin loop structure were ligated to prepare for hybridization. The library fragments were purified with the AMPure XP system (Beckman Coulter, Beverly, USA) for selecting cDNA fragments of preferentially 250~300 bp in length. Then USER Enzyme (NEB, USA) was used, adaptor-ligated cDNA at 37 °C for 15 min, followed by 5 min at 95 °C. PCR was then performed with Phusion High-Fidelity DNA polymerase, Universal PCR primers, and Index (X) Primer. Finally, PCR products were purified (AMPure XP system) and the quality of libraries was assessed by the Agilent Bioanalyzer 2100 system.

Index-coded samples were clustered on the cBot Cluster Generation System using TruSeq PE Cluster Kit v3-cBot-HS (Illumina) following the manufacturer’s instructions. After clustering generation, the library preparations were sequenced on the Illumina Novaseq 6000 platform to generate 150-bp paired-end reads.

Identification of Differentially Expressed Genes and Functional Enrichment Analysis

Clean data were filtered using trimmomatic (Bolger et al. 2014) to remove reads that contained adapter contamination, low-quality nucleotides, and unrecognizable nucleotides (N) from the raw data. The genome and annotation files of O. minor were downloaded from the GigaScience Database (http://gigadb.org/dataset/100503). HISAT2 v2.1.0 was used to build reference genome indexes (Kim et al. 2015), and then the clean data were aligned to the O. minor genome. FeatureCounts was applied to quantify gene expression (Liao et al. 2014). Differential expression analysis was performed by DESeq2 (Love et al. 2014). Differentially expressed genes (DEGs) were identified as those with |log2(FoldChange)|> 1 and adjusted P-value < 0.05. To functionally annotate DEGs, all genes were first annotated using eggnog-mapper v2 (Cantalapiedra et al. 2021). Then the R package AnnotationForge (https://bioconductor.org/packages/AnnotationForge/) and clusterProfiler (Wu et al. 2021) were used to conduct GO and KEGG enrichment analysis.

Validation of RNA-seq Data by qRT-PCR

Twelve differentially expressed genes related to gonadal development were validated by qRT-PCR. All primers are shown in Table S1, with β-actin selected as the internal control. To synthesize cDNA, the remaining RNA from 12 gonad samples after RNA-seq sequencing was used with PrimeScript RT reagent Kit (Takara, Dalian, China) according to the manufacturer’s instructions. Then, qRT-PCR was performed on the LightCycler 480 real-time PCR instrument (Roche Diagnostics, Burgess Hill, UK) by QuantiNova SYBR Green PCR Kit (QIAGEN, Germany) in 10 μl reactions. Each reaction contained 1 μl of cDNA, 5 μl of 2 × SYBR Green PCR Master Mix, 2.6 μl of ddH2O, 0.7 μl of forward primer, and 0.7 μl of reverse primer. The cycling parameters were as follows: 95 °C for 5 min, followed by 40 cycles of 95 °C for 5 s, and 60 °C for 30 s. The relative expression levels were calculated by CT methods (Schmittgen and Livak 2008).

Results

Annual Changes of TW and GSI

Changes in TW and GSI from September 2020 to June 2021 are shown in Fig. 1. The TW trend was consistent in both females and males and was comparable to the letter “M”. There were two growing periods, one from September to December and the other from March to April (Fig. 1A). The GSI values of females showed an increasing trend, with the minimum value occurring in September and the maximum value in June of the following year. While GSI values in males fluctuated at around 3% (Fig. 1B).

Fig. 1
figure 1

Morphological measurements and macroscopic maturity stages of O. minor in the male and female reproductive system. A Monthly change in the total body weight. B Monthly changes in gonad somatic index. C Immature ovary. D Mature ovary. E Maturing testis. F Mature testis. G Spent testis. Scale bar: C–G = 10 mm

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The macroscopic maturity stages of females and males are presented in Fig. 1C–G. During the immature period, the female reproductive system was small and translucent with indistinct egg grains (Fig. 1C). However, in the mature stage, the female reproductive system appeared yellowish and the ovaries were significantly enlarged and fully developed. The egg grains could be observed with the naked eye, and the oviducal glands were white (Fig. 1D). Three macroscopic stages of testicular development were observed, including maturing, mature, and spent. In the maturing stage, spermatophores were presented in males (Fig. 1E). During the mature stage, two to seven spermatophores were observed (Fig. 1F). At the spent stage, the male reproductive tract was flaccid and the testes were cream yellow with 2–3 spermatophores (Fig. 1G).

Oogenesis

According to microscopic observation, we identified 9 oocyte stages and described the characteristics as follows.

  • Stage 1 — Oogonia (OO): Small round cells of 11.3 to 26.9 μm diameter, with almost no cytoplasm. These cells are near the germinal cords (Fig. 2A).

  • Stage 2 — Early primary oocyte (EPO): Oocytes are oval-shaped, binding to several follicles. Oocytes range from 33.8 to 74.7 μm in diameter (Fig. 2B).

  • Stage 3 — Late primary oocyte (LPO): Oocytes are surrounded by a layer of small follicle cells. The diameters of oocytes vary between 97.1 and 194.0 µm. The cytoplasmic portion of the cell is larger than the nucleus. Lipid inclusions can be observed in the cytoplasm (Fig. 2C).

  • Stage 4 — Previtellogenic oocyte (PVO): The follicle cells proliferate intensively. The follicle epithelium consists of two layers of follicular cells, with the inner layer of follicle cells forming folds and penetrating deep inside the cell. The diameter of the oocyte increases (179.6 to 394.2 µm). The nucleoli begin to degenerate, and the yolk globules are initially produced. The lipid inclusions multiply and enlarge (Fig. 2D).

  • Stage 5 — Vitellogenic oocyte (VO): Oocyte size increases greatly, with the diameters ranging from 235.3 to 735.2 µm. The follicular epithelium is active during vitellogenesis and chorion formation. Follicular folds migrate to the periphery of the oocyte through yolk formation (Fig. 2E).

  • Stage 6 — Advanced vitellogenic oocyte (AVO): Oocytes reach the maximum size (1246.1 to 3144.7 µm). The cytoplasm is filled with yolk granules and surrounded by the chorion (Fig. 2F).

  • Stage 7 — Ripe oocyte (RO): Oocytes are released by the preovulatory follicle. The cytoplasm contains yolk granules and the folds are completely reabsorbed, involved, and protected by the chorion. These oocytes are ready for ovulation. The oocyte diameters varied between 2187.1 and 3405.9 µm (Fig. 1G).

  • Stage 8 — Post-ovulatory follicle (POF): The post-ovulatory follicle is generated by the follicular epithelium after ovulation. Follicles are irregularly shaped and contain a star-shaped lumen filled with fibrous material and highly amorphous and basophilic bodies (Fig. 2H).

  • Stage 9 — Atretic oocyte (AO): The follicular epithelium in atretic oocytes is disorganized. The fibrillar connective tissue is replaced by collagen fibers. The chorion is disorganized and fragmented (Fig. 2I).

Fig. 2
figure 2

Oogenesis of ovaries in O. minor. A Oogonia (OO). B Early primary oocyte (EPO). C Late primary oocyte (LPO). D Previtellogenic oocyte (PVO). E Vitellogenic oocyte (VO). F Advanced vitellogenic oocyte (AVO). G Ripe oocyte (RO). H Post-ovulatory follicle (POF). I Atretic oocytes (AO). Scale bar: A–C, H, I = 20 μm; D, E = 50 μm; F, G = 200 μm

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Four types of oocytes (OO, EPO, LPO, and RO) typically appeared in June. In September, three types of oocytes (PVO, POF, and AO) could be observed. The VO phase was the primary type of oocyte that existed from September to March of the subsequent year, while AVO mainly occurred from April to May.

Spermatogenesis

Four microscopic stages of testicular development were observed and described below.

  • Stage 2a (developing): In the seminiferous tubules, spermatogonia, a large number of primary and secondary spermatocytes, and a small amount of sperm cells are observed, with the absence of spermatozoa (Fig. 3A).

  • Stage 2b (maturing): The seminiferous tubules are clearly defined with the presence of spermatogonia, primary and secondary spermatocytes, and spermatids. Spermatozoa are visible in all seminiferous tubules (Fig. 3B).

  • Stage 3a (mature): The seminiferous tubules are large and well-defined with no intercellular gap. All cell types are present, and there is an abundance of spermatozoa in the central lumen (Fig. 3C).

  • Stage 3b (spent): Only a limited number of primary and secondary spermatocytes, spermatids, and spermatozoa are scattered throughout the seminiferous tubules (Fig. 3D).

Fig. 3
figure 3

Spermatogenesis of testes in O. minor. A Developing. B Maturing. C Mature. D Spent. Spermatogonia (SPG), primary spermatocytes (SPC I), secondary spermatocytes (SPC II), spermatids (SPD), spermatozoa (SPZ), and flagella (F). Scale bar: A–C = 20 μm; D = 100 μm

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The developing stage of testes was observed in an octopus weighing 27 g in June, whereas the maturing stage was mainly examined in September. The mature stage was the period that lasted the longest from September to March of the following year. The spent phase primarily occurred from April to June.

Data Quality Control and Alignment

A total of 528,428,198 raw reads were obtained from 12 gonad samples, which were divided into 4 groups: AVO, VO, Spent, and Mature. After filtering, 505,726,548 clean reads were retained. The values of Q20 (%) and Q30(%) exceeded 98.22% and 93.68% (Table S2), respectively. Clean reads were aligned to the O. minor genome, with overall alignment rates ranging from 76.8 to 89.9%. All raw data were submitted to the Sequence Read Archive (SRA) of the National Center for Biotechnology Information (NCBI) under the Bioproject PRJNA993392, with accession numbers SRR25224769-SRR25224780.

Differential Expression, GO, and KEGG Enrichment Analysis

The counts obtained from FeatureCounts were utilized for differential expression analysis. In the “AVO vs VO” group, 1095 DEGs showed significantly differential expression, including 534 upregulated genes and 561 downregulated genes (Fig. 4A, B). Meanwhile, in the “Spent vs Mature” group, a total of 2468 DEGs were significantly differentially expressed, consisting of 909 upregulated genes and 1559 downregulated genes (Fig. 4A, B). Figure 4C displays the gene expression profiles of 10,175 DEGs in four groups, revealing great differences between the ovary and testis.

Fig. 4
figure 4

DEGs of ovaries and testis in O. minor. A Upregulated and downregulated DEGs in the ovary and testis. B Venn diagram of DEGs. C Heat map of 10,175 DEGs

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To explore the potential regulatory function and pathway of DEGs in gonad development, we performed the enrichment analysis. The results showed that 126 GO terms were significantly enriched in the group “AVO vs VO” (Table S3), mainly associated with biological processes such as “Cytoplasmic translation”, “RNA splicing”, and “G1/S transition of the mitotic cell cycle” (Fig. 5A). In the KEGG analysis, five pathways were found to be significantly enriched and downregulated, namely “Ribosome”, “Cell cycle-yeast”, “Cell cycle”, “Progesterone-mediated oocyte maturation”, and ''Meiosis-yeast" (Fig. 5B, Table 1), suggesting a weakened yolk deposition activity during the further maturation of the oocytes.

Fig. 5
figure 5

Enrichment analysis of DEGs in ovaries of O. minor. A Gene ontology (GO) annotation. B KEGG pathway of DEGs. Asterisks represent significantly enriched terms

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The GO enrichment analysis presented that 674 GO terms of 2468 DEGs were considerably enriched in “Spent vs Mature” (Table S4). These terms primarily centered on “Meiosis”, “RNA splicing”, and “Regulation of cyclin-dependent protein serine/threonine kinase activity” (Fig. 6A). Thirteen KEGG pathways were significantly enriched, consisting of four upregulated DEGs pathways and nine downregulated DEGs pathways (Table 1), respectively. The pathways “Spliceosome”, “mRNA surveillance pathway”, “Nucleocytoplasmic transport”, and “Protein processing in endoplasmic reticulum” were observed to be upregulated. Meanwhile, “Oxidative phosphorylation”, “Carbon metabolism”, and “Glycolysis/gluconeogenesis” pathways were downregulated (Fig. 6B).

Fig. 6
figure 6

Enrichment analysis of DEGs in testes of O. minor. A Gene ontology (GO) annotation. B KEGG pathway. Asterisks represent significantly enriched terms

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Table 1 Significantly enriched KEGG pathways in “AVO vs VO” and “Spent vs Mature”
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Validation of RNA-seq Data by Quantitative Real-Time PCR

To verify the reliability of the RNA-seq data, we selected 12 genes for qRT-PCR validation, including 7 DEGs for ovaries and 5 DEGs for testes independently. Overall, the qRT-PCR results were consistent with the RNA-seq data (Fig. 7).

Fig. 7
figure 7

Validation of differentially expressed genes in O. minor. A Seven DEGs in group “AVO vs VO”. B Five DGEs in “Spent vs Mature”

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Discussion

O. minor is an economically valuable species in northern China, but its gonadal research is currently limited. Identifying and categorizing stages of maturity is essential for artificial cultivation and fisheries. Therefore, this study investigated the histological processes of gonad development and changes in BW and GSI of both female and male O. minor. Additionally, we selected two key histological stages in the ovaries and testes of O. minor to conduct transcriptome analysis and identify potentially critical regulatory pathways and genes.

Reproductive Strategy and Histology of Gonadal Development

Like most cephalopods, O. minor is a short-lived and rapidly developing species. In females, TW and GSI increased rapidly from September to December, then decreased and remained flat until March of the subsequent year. Proteins in muscle serve as the main energy reserve for cephalopods, while lipids are critical energy sources for the ovary (Rosa et al. 2004, 2005). In O. vulgaris, Sieiro et al. (2020) recorded a notable decline in lipids and an increase in proteins in the digestive glands of females from winter to spring, but no significant changes were observed in these two biochemical compositions in the mantle. It has been demonstrated that proteins and lipids in the mantle and digestive glands of cephalopods decrease during the period of spawning and post-spawning (Kilada and Riad 2008; Morillo-Velarde et al. 2013). However, no relevant studies have been conducted on O. minor. We hypothesize that females may initially use the energy obtained from food to maintain the basic physiological metabolism, and possibly consume lipids from the digestive glands to provide energy during the overwintering period, which may explain the observed decrease and stagnation in both BW and GSI. Yolk deposition in cephalopods is a long process. Oocytes in the VO phase require around 6 months to progress to the AVO stage due to the insufficient lipid supply during the overwintering period. When temperatures rise in March, abundant food will lead to weight gain and make the ovary fuller until ovulation. In some cephalopods, females would limit the growth of somatic cells to meet reproductive development, while males may experience concurrent somatic growth with reproductive cell development (Jackson et al. 2004; Lin et al. 2015). Therefore, we speculate that this reproductive strategy also applies to O. minor, which may account for the higher mean BW of the males compared to the females collected simultaneously in this study.

Changes in the female reproductive system during maturation were evident. Figure 2C, D show that ovaries are much fuller and light-yellowish, and the oviducal glands significantly enlarged with the color changed from translucent to creamy white. However, this color transition of oviducal glands is different from the brown or amber color found in many other mature octopod species (de Lima et al. 2013; Olivares et al. 2017). The peripheral gland of the female reproductive system was reported to produce mucins as the ovary matures, which in turn enlarges the oviducal gland (Froesch and Marthy 1975). Additionally, we also noticed that females were generally smaller than males, probably because they allocated more energy to reproduction.

In cephalopods, it is very common that males undergo a very short sub-mature period and mature much earlier than females, with spermatophores present for most time of the year (Avila-Poveda et al. 2009; López-Peraza et al. 2013; Otero et al. 2007). Females can store sperm in oviducal glands for months (Mangold 1987), enabling males to mate with females almost all year round. In this study, we observed that the number of spermatophores ranged from 2 to 9 in male O. minor, which was much fewer than that of octopod species with a benthic stage like O. vulgaris. Besides, we found that the male O. minor with spermatophores weighed 32 g, indicating that juveniles are capable of mating.

The histological process of gonad development in O. minor is similar to other cephalopods. According to the histological classification of O. vulgaris (ICES 2010), nine and four stages were observed in females and males, respectively. Oocyte development in O. minor is asynchronous, as oocytes closer to the germinal cord mature more slowly. The appearance of the previtellogenic oocyte signifies the beginning of maturation and yolk deposition, which lasts for at least 6 months in O. minor. In April, spermatozoa were found in the oviducal gland of O. minor, whereas no spermatozoa were observed before this time, suggesting that the peak time of the mating may occur in April. The process of spermatogenesis in O. minor is akin to that of O. vulgaris and A. fangsiao (Cuccu et al. 2013; Wang et al. 2015). In the seminiferous tubules, the outer primary spermatocytes undergo meiosis and mitosis to produce a large number of spermatozoa in the center of the tubular lumen. As mentioned above, males matured very fast and spermatophores were observed in all 10 months of sampling from 2020 to 2021, which was in accordance with the result of other studies (Avila-Poveda et al. 2009; Cuccu et al. 2013; de Lima et al. 2013).

O. minor is known to spawn from July to late August (Liu 2013; Qian 2011) and incubate for 72 to 89 days under temperatures of 21 to 25 ℃ (Qian et al. 2013). Consequently, it seems reasonable to assume that O. minor hatchlings appear in October. However, the majority of octopuses collected in September weighed about 50 g, and the others weighed around 110 g. Previous data from artificial culture showed that O. minor larvae needed about 200 days to reach a weight of 50 g and 250 days to achieve 110 g under adequate food conditions (Zheng et al. 2014). However, these two results seem contradictory if the embryo development time in the wild coincides with that under artificial conditions. So, O. minor in Swan Lake was investigated in June 2023. As a result, eleven juveniles of 1.2 to 2.0 g were successfully captured and identified as O. minor by DNA barcoding. Research has indicated that an artificially hatched O. minor juvenile weighed 1.33 g at 75 days (Nan 2020). Therefore, it is highly likely that the juveniles in Swan Lake hatched around March. In addition to this, we also collected a larger O. minor weighing 27 g.

Compared to shallow waters, deep-sea water temperatures are lower. The incubation period of some deep-sea cephalopod eggs has been reported to be several years (Robison et al. 2014; Wood JB et al. 1998). Therefore, based on the current results and analysis, we hereby propose that O. minor has a lifespan of at least 18 months and may employ a special reproductive strategy with both egg and larval overwintering periods. In northern China, seawater temperatures gradually decrease from September and can even drop to 0 ℃ in December. The biology zero of fertilized eggs for O. minor was 3.79 ℃, indicating that O. minor could well adapt to low temperatures (Liu 2013). To make the larvae survive, O. minor may adopt an egg overwintering strategy; that is, fertilized eggs laid in July to August develop slowly at low temperatures and then hatch in March of the following year when the seawater temperature is suitable. After half a year, larvae would grow to 50 g by September. On the other hand, the presence of the larger juvenile suggests that O. minor is likely to have a larval overwintering period as well. Briefly, fertilized eggs hatch from October to November and grow slowly at low temperatures. However, there is no observational study of O. minor fertilized eggs hatching in the wild, and our speculations need to be confirmed by further field research.

Key Pathways of Ovarian Development in O. minor

Yolk deposition of oogenesis in cephalopod is a critical and long process that typically lasts for several months (de Lima et al. 2013; Rodríguez-Rúa et al. 2005; Wang et al. 2015). As shown in Table 1, five pathways are significantly enriched in group “AVO vs VO”. A total of 75 downregulated genes were clustered in the ribosomal, cell cycle, progesterone-mediated oocyte maturation, and meiotic pathways that were highly associated with oocyte development.

Yolk in oocytes is mainly synthesized by ribosomes in follicular cells and subsequently transported to the cytoplasm of oocytes. As the oocyte matures, yolk synthesis activity gradually decreases. The KEGG result indicated that twenty-three ribosomal protein genes or rRNA molecules were enriched in the ribosomal pathway, suggesting that these RPLs and RPSs are involved in the biosynthesis of oocyte yolk, which is common in other studies (Rojo-Bartolome et al. 2016; Sun et al. 2022). The histological results showed that the follicular epithelium, composed of follicular cells, was highly active during the VO stage. As the oocytes developed into AVO, the cytoplasm was filled with yolk granules, and the ability to synthesize ribosomes was diminished.

Animals complete the first meiotic division before ovulation, in which the transition from G2 to M phase is one of the key processes in oocyte maturation. CPEB controls mRNA polyadenylation during oocyte maturation in the progesterone-mediated oocyte maturation pathway (Stebbins-Boaz et al. 1996), and it also increases the mitosis-promoting factor (MPF) Cdc2/cyclin B by downregulating Myt1 (Nakajo et al. 2000). Additionally, the role of progesterone-mediated oocyte maturation in many aquatic animals is closely related to the cell cycle signaling pathway (Jia et al. 2018). In the present study, GO term cyclin-dependent protein kinase holoenzyme complex (GO: 0043073) was notably enriched, and CDK1, CDC25A, and CCNB2 genes were upregulated in the VO stage and were co-regulated in both “Progesterone-mediated oocyte maturation” and “Cell cycle” pathways. In the “Cell cycle” pathway, CDC25 mediates dephosphorylation to activate the CyclinB-CDK1 complex, which phosphorylates several substrates and triggers centromere segregation and chromosome condensation. Once chromosomes are condensed and aligned at the midplate, CDK1 activity will be turned off by WEE1- and PKMYT1-mediated phosphorylation, allowing for sister chromatid separation, chromosome decondensation, reorganization of the nuclear membrane, and cytoplasmic division (Morgan 1995; Mueller et al. 1995). Our results showed that these three genes were upregulated during the VO stage, highlighting their indispensability in the yolk synthesis of O. minor, and suggesting that oogenesis involves a complex process of multiple factors and pathways. Also, MCM4, MCM7, BUB1, and SMC1B were upregulated in the AVO stage, indicating that they are also relevant to the cell cycle and meiosis.

Key Pathways of Testicular Development in O. minor

Spermatogenesis comprises three stages, including spermatogonia mitosis, spermatocyte meiosis, and spermatocyte metamorphosis into spermatozoa. Testes classification depends on the presence and proportion of primary spermatocytes, secondary spermatocytes, spermatocytes, and spermatozoa. In the present study, the testis exhibited a more complicated developmental process, as more KEGG pathways were significantly enriched in the testes than in the ovaries.

Energy metabolism in the testis is essential for spermatozoa. In this study, we observed the significant downregulation of “Oxidative phosphorylation”, “Carbon metabolism”, and “Glycolysis/gluconeogenesis” pathways, which implies that the process of testis recession may be regulated by reduced energy metabolism and decreased energy supply. The “Oxidative phosphorylation” pathway is also critical for testis as it is involved in several processes, such as sperm survival, development, and motility, by functioning in the mitochondria of germ cells and supporting cells, thus affecting sperm quantity and quality (Wang et al. 2022). Like the oriental river prawn Macrobrachium nipponense (Jin et al. 2022), “Oxidative phosphorylation” was the most significantly enriched pathway in O. minor testes. ATP synthase subunits participate in the oxidative phosphorylation process of energy production in sperm mitochondria, which was found to be crucial in the formation of Drosophila sperm mitochondria (Sawyer et al. 2017). In this study, several ATP synthase subunit genes such as ATP5L, ATP6V0C, and ATP5F1 were downregulated, signifying that these genes may decrease the level of energy metabolism in spermatozoa, in turn, affect sperm motility and survival. To compensate for the deficiency of oxidative phosphorylation, the glycolytic pathway can provide an adequate energy supply. Glycolysis promotes the conversion of glucose (C6H12O6) to pyruvate (CH3COCOO− + H+), releasing free energy to form the high-energy molecules ATP, and reducing nicotinamide adenine dinucleotide (Lubert 1995). Gluconeogenesis, which synthesizes glucose from non-carbohydrate precursors, is an important source when the glycolytic response is insufficient (Gerich 2010). PGAM1 is one of the key enzymes in the glycolytic pathway and studies have suggested that it may be related to controlling the development of spermatogenic cells (Rato et al. 2012, 2016). We also noticed other main genes of the glycolytic pathway such as ENO2, GAPDH, and GPI were significantly downregulated in the Spent phase, suggesting that the glycolytic pathway of these genes acts in synergy with oxidative phosphorylation to provide energy for testis development during the reproductive season.

In this study, we found that several pathways involved in cellular repair, such as “mRNA surveillance pathway”, “Spliceosome”, “Nucleoplasmic transport”, and “Protein processing in endoplasmic reticulum”, were significantly upregulated. Cell repair plays a crucial role in the restoration and protection of spermatozoa during spermatogenesis (Gunes et al. 2015). Splicing, an essential regulatory process for gene expression, involves the removal of unrelated or incorrect introns from pre-mRNA, resulting in the generation of multiple mRNA isoforms, and leading to a diverse of gene functions (Nilsen 2003; Will and Luhrmann 2011). The mRNA surveillance pathway maintains precise control over intracellular signaling and metabolism by monitoring processes such as mRNA transcription, splicing, and stability. Previous studies have demonstrated the significance of mRNA surveillance and spliceosome pathways in the maturation of spermatocytes in aquatic animals (Jin et al. 2020; Xu et al. 2015). The nucleocytoplasmic transport and protein processing in the endoplasmic reticulum signaling pathways could cooperate with various pathways, including cell cycle regulation and apoptotic response (Ferrando-May 2005; Major et al. 2011; Naidoo 2009). As an important transcription factor in the endoplasmic reticulum stress pathway, ATF6 promotes many processes, such as endoplasmic reticulum protein folding and subsequent secretion by regulating promoter region activity and inducing cellular stress response (Wang et al. 2000). A study demonstrated that knocking out ATF6 in mice resulted in decreased male fertility (Yu et al. 2022). Furthermore, silencing ATF6 promotes the autophagic pathway and affects apoptosis (Liu et al. 2020). Thus, the upregulation of the ATF6 might be associated with the reparation of declining spermatozoa and the maintenance of homeostasis. HSP90 is a molecular chaperone that takes part in numerous cellular processes. Research has shown that knocking out the Hsp90α gene leads to a significant reduction in high steady-state levels of HIF-1α in the testis, preventing sperm production and causing sterility in mice (Tang et al. 2021). On the other hand, HSP90 could also enhance glycolysis and proliferation and reduce apoptosis (Xu et al. 2017). Thus, the upregulation of HSP90 highlighted its significant function in maintaining spermatozoa homeostasis during the spent phase of male O. minor. In a word, the upregulated signaling pathways collaborated to enhance cellular repair functions and maintain the homeostasis of O. minor testis.

Conclusion

In this study, we provided integrated data on the morphology, histology, and transcriptome of O. minor. We proposed that O. minor had a lifespan of at least 18 months and had both eggs and larval overwintering periods. Pathways identified in two stages of ovarian development were primarily linked to the “Ribosomal”, “Cell cycle”, and “Progesterone-mediated oocyte maturation” pathways. Genes such as CDK1, CDC25A, and CCNB2 were considered to play key roles in yolk deposition. Energy metabolic pathways, such as “Oxidative phosphorylation” and “Glycolysis/gluconeogenesis”, may contribute to energy supply for male reproduction. Relevant genes, including those in the ATP synthase subunit family and PGAM1, may be involved in mediating spermatogenesis. Signaling pathways related to cell repair were important for maintaining homeostasis in the recessionary testis of O. minor. Our study provides the molecular basis for the artificial breeding and resource conservation of O. minor.