Effect of acid modification of Biochar derived from spent mushroom substrate on the production of oyster mushroom (Pleurotus ostreatus)

Abstract

Spent mushroom substrate (SMS) disposal presents significant environmental challenges in the edible fungi industry. Converting SMS into biochar offers a promising resource utilization approach; however, the original biochar properties limit its effectiveness in mushroom cultivation applications. In this study, SMS biochar was modified by phosphoric acid treatment to form acid-modified mushroom substrates (AMMS), and the influence of acid-modified mushroom substrates on the production of oyster mushroom was evaluated. According to the obtained data, Acid modification significantly increased the surface area and micropore volumes of biochar. Fourier transform infrared spectroscopy (FTIR) analysis revealed the hydrophilicity and adsorption capacity of acid-modified biochar have been enhanced. The highest-performing treatments in AMMS showed 13–16% yield increases compared to unmodified biochar treatments, with harvest time advanced by 2.5 days. The safety assessment confirmed that all treatments with biochar added had no video security risks. In conclusion, phosphoric acid modification of SMS biochar significantly improves its performance as a mushroom cultivation substrate additive. This approach provides an effective strategy for SMS resource utilization and offers technical support for optimizing oyster mushroom cultivation practices.

Introduction

Oyster mushroom (Pleurotus ostreatus) belongs to the family of edible fungi, which is extensively cultivated and consumed around the world. It is highly favored for its unique flavor, rich nutritional value and potential medicinal properties^1,2. However, after the harvest of edible fungi, about five kilograms of SMS is produced per one kilogram produced mushroom³. SMS is rich in organic matter, residual mycelium and various enzymes. However, its direct disposal (such as piling up or landfilling) not only occupies land resources but also leads to the secondary pollution of the environment as a result of organic matter decomposition including the generation of leachate and greenhouse gases⁴. Therefore, how to handle and utilize SMS efficiently and environmentally friendly has become an important issue for the sustainable development of the edible fungi industry. At present, the reuse approaches of SMS include being used as soil conditioners, organic fertilizers, animal feed, raw materials for bioenergy production, etc^5–7. Biochar is characterized by high carbon content, porosity and large specific surface area. Incorporating biochar into the substrate for edible fungi can enhance its water and nutrient retention capacity. Meanwhile, the large specific surface area provides more attachment points for mycelium, thereby promoting the growth of mushroom. Therefore, converting SMS into biochar is a highly promising way of resource utilization.

Biochar is a category of carbon-rich porous materials manufactured by the pyrolysis of biomass under anaerobic or oxygen-restricted conditions at elevated temperatures⁸. Using SMS as raw material to prepare biochar can not only effectively deal with waste but also produce products with high added value. The application of biochar in the cultivation substrate of edible fungi has been an emerging research direction in recent years. Studies have shown that introducing optimum amounts of biochar to the cultivation substrate is able to enhance its physical properties, such as enhancing water retention and aeration, regulating pH value, adsorbing and slowly releasing nutrients, and even growth inhibition of some pathogenic bacteria, thereby promoting mycelial growth and increasing the yield and quality of mushrooms³. However, some properties of the original biochar (such as the types and quantities of surface functional moieties, pore size distribution, pH value, etc.) may not be optimal for some applications.

To overcome this shortcoming of biochar, it is usually desired to modify it. Common biochar modification approaches are physical modification (such as steam activation, ball milling), chemical modification (such as acid and alkali treatment, oxidant treatment, metal salt impregnation), and biological modification^9–11. Among them, acid modification is a commonly used and effective chemical modification method. The physicochemical properties of biochar can be significantly altered by treating it with inorganic or organic acids^12,13.

The acid modification of biochar mainly affects the following aspects: Firstly, acid treatment removes the ash and impurities clogged in the channels of biochar, etches the pore walls, thereby forming new pores or expanding the existing ones, increasing the total pore volume and specific surface area of biochar, especially the ratio of micropores to meshes^8,14. Secondly, acid modification can introduce or increase oxygen-containing functional moieties, like carboxyl groups (-COOH), hydroxyl groups (-OH), and phenolic hydroxyl groups, on its surface^15,16. The increase of these functional groups is able to enhance the polarity and hydrophilicity of the biochar surface, strengthen its interaction with nutrient ions (e.g. complexation, ion exchange, and hydrogen bonding), thereby improving its ability to serve as a nutrient sustained-release carrier¹⁷ (Fig. 1). Finally, pickling can effectively remove part of the ash in biochar (mainly soluble mineral salts), increase the carbon content, and thereby improve the purity and stability of biochar¹⁸. These changes are all conducive to providing favorable conditions for the growth of edible mushrooms.

Fig. 1.

Acid-modified biochar enhances the availability of nutrient ions in mushroom cultivation substrates.

Although the application of biochar in mushroom cultivation^19,20acid-modified biochar in adsorption and soil improvement^12,21regarding the application of acid-modified SMS biochar in oyster mushroom cultivation, and its effects on substrate characteristics have been studied previously, studies about the mycelial growth, mushroom yield, quality, economic benefits and product safety in this field are still rare. The previous work of this research team found that adding a certain amount of SMS biochar to the mushroom substrate promotes mycelial growth, enhances the yield of Pleurotus ostreatus and truncates the cultivation duration³. On this basis, this study intends to modify SMS biochar using phosphoric acid, aiming to further optimize its performance and explore its effect as an additive for the cultivation substrate of oyster mushroom. This research aims to provide a new strategy for the efficient utilization of SMS, and offer scientific basis and technical support for optimizing the cultivation techniques of oyster mushroom, improving production efficiency and ensuring product safety.

Materials & methods

Preparation of acid-modified Biochar

The production of biochar from SMS is described in our previous study³. The SMS in this study were collected from mushroom cultivation in Chongqing Academy of Agricultural Sciences, Chongqing, China. The SMS was collected after the mushroom harvest, and stored under dry conditions before it was converted to biochar. Briefly, the SMS was placed in an oven at 85 °C for six hours, and then the dry samples were grounded to pass a 1-mm sieve. The samples were then loaded into a quartz tube furnace containing a quartz crucible, which was connected to a dinitrogen gas (RISING GAS, Purity 99%) supply device to create oxygen-free conditions throughout the pyrolysis process. The temperature in the tubular furnace was set at 500 °C at an increasing rate of 25 °C min^− 1. The reaction pressure was 50–60 kPa. The pyrolysis continued for three hours. The newly produced biochar were removed out of the tubular furnace after they had cooled to room temperature. Acid-modified SMS biochar was prepared by thoroughly mixing 300 mL of 3.0 mol·L⁻¹ H₃PO₄ with 20 g of biochar, followed by impregnation, stirring for 1 h and shaking for 12 h.

Experimental design

The Pleurotus ostreatus strain used in the experiment was acquired from the Beijing Institute of Agricultural Resources and Regional Planning, Chinese Academy of Agricultural Sciences. The biochar-based substrates were prepared by mixing the cultivation ingredients with the acid-modified biochar and biochar to form the acid-modified biochar mushroom substrates (AMMS) and biochar mushroom substrates (BMS), respectively. The names and quantities of additional ingredients as well as mushroom baglogs and the details of the mushroom cultivation process are mentioned in our previous publication²². Herein, seven treatments are introduced: CK (no biochar), B1 (5 wt% biochar), B2 (10 wt% biochar), B3 (15 wt% biochar), PB1 (5 wt% phosphoric acid-modified biochar), PB2 (10 wt% phosphoric acid-modified biochar), and PB3 (15 wt% phosphoric acid-modified biochar). Cottonseed husks, wheat bran, and the other main components were thoroughly mixed with biochar, then the moisture was adjusted to 60%, and the substrates were packed in 18 × 45 cm polyethylene plastic bags, which were termed mushroom baglogs. Each bag was filled with approximately 1.3 kg substrate and sterilized at 105 °C for 10 h. After the baglogs were autoclaved and cooled to room temperature, oyster mushroom spawn was inoculated into the baglogs, and the baglogs were stored at a mushroom house with an intelligent monitoring system. Each treatment had three replicates and there were 30 mushroom baglogs per replicate.

Determination of mushroom substrate properties

Compositions and elemental of AMMS and BMS were determined using thermogravimetric analyser (Mettler-Toledo, Switzerland)²³ and elemental (Elementar Analysen Systeme GmbH, Germany) analyser²⁴. The specific surface area of each treatment was determined using an automatic nitrogen-specific surface area analyser (Micromeritics, USA), and then calculated by the Brunauer, Emmett, and Teller (BET) equation.

Determination of substrate moisture, EC and pH during mushroom growth

The characteristics of the mushroom substrate were measured four days after spawning when the primordium appeared. The data were recorded again before the second harvest. The pH and EC of the biochar samples were measured directly in one-fifth water suspension solutions with a pH meter (INESA Scientific Instrument Co., Ltd., China). The moisture was measured using a soil humidity sensor (Jianda Instrument Co., Ltd., China).

FTIR spectra measurements

FTIR spectra were acquired from pellets consisting of 1–2 mg of samples and 100 mg KBr, spanning the range of 700–4000/cm. Each sample underwent 20 scans, and all spectra were subsequently normalized.

Measurement of mycelium growth and mushroom yield

The growth of mushroom mycelia was quantified by the area of substrate colonized. Measurements were taken every 4 days post-spawning until 28 days. Following primordium formation, the cover of the mushroom bag was removed to enable fruit body development, and mushroom yields were recorded for two harvesting cycles.

Food safety risk analysis

Mushrooms grown on different substrates were harvested and kept frozen in liquid nitrogen until the metabolites and contaminant analysis. The freeze-dried sample was ground, and then extracted with a mixture of 5 mL 50% methanol and 5 mL chloroform. The mixture was centrifuged, and the lower layer of organic chloroform was collected into a glass bottle, dried completely with a high-speed vacuum, and stored at -80 °C. Prior to nuclear magnetic resonance (NMR) analysis using a Bruker DRX-400 advance (400 MHz) spectrometer (Bruker, Germany), the extracted metabolites were dissolved in 600 mL 99.8% chloroform-d and transferred into a 6 mm NMR tube for organic metabolite detection. NMR samples were collected according to a previous study²⁵. Concentrations of contaminants in mushrooms were determined using an atomic absorption spectrophotometer (PerkinElmer, Inc., USA) following the manufacturer’s procedure²⁶.

Statistical analysis

The data in the tables and figures are expressed as the means of all replicates ± standard deviation (SD). Data were subjected to analysis of variance using the SAS version 9.2 (SAS Institute, Cary, NC). Means were compared using the least-significant difference (LSD) test. Differences were declared significant at P < 0.05. All figures were generated using the Origin 2019 Pro.

Results and discussion

The properties of AMMS

Acid modification considerably changed the ratio of the constituting contents of the substrate (Table 1). With the increase of the proportion of biochar in the substrate, the amount of volatile matter reduces while the fixed carbon ratio increases in each treatment. Compared with BMS, the fixed carbon content of AMMS has further increased. The ash and water contents of each substrate treated with biochar were lower than those of CK without it. Except for CK, other samples showed no dramatic differences in their ash and water ratios. In terms of elements, compared with CK, the carbon and oxygen contents enhanced and decreased, respectively, in BMS and AMMS treatments. The O/C value increase in the AMMS treatment indicates that the hydrophilicity of the modified biochar mushroom substrate was significantly enhanced. During the pyrolysis process, due to the combined influence of oxidation reactions and high-temperature pyrolysis, biochar often exhibits smaller volatile matter and larger fixed carbon ratios. The phenomenon of reduced volatile matter and increased fixed carbon observed in this study is very likely closely related to the reconstruction of organic components and the evaporation of volatile materials during the pyrolysis process²⁷. Meanwhile, the acid modification process can further promote the rearrangement of functional groups on the surface of biochar, forming a stable carbon skeleton, thereby further increasing the fixed carbon content^28,29. It is particularly noteworthy that in the acid-modified substrate, although the overall oxygen content is low, the O/C ratio rises, which indicates that despite the removal and transformation of some non-polar components during the acid treatment, some oxygen-containing moieties (e.g. hydroxyl and carboxyl) may be retained or reintroduced in a higher proportion, thereby improving the hydrophilicity of the substrate^30,31. This is of great significance for mushroom culture substrate, because the hydrophilicity of substrate not only affects the uniform distribution of water and nutrients but also may further regulate the activity of the growth of mushroom mycelium³².

FTIR analyses of Biochar

Acid modification can endow biochar with more oxygen-containing moieties, improving the hydrophilicity of the mushroom substrate while rising its adsorption capacity for nutrient ions³³. Compared with biochar (BC), the intensity of the -OH stretching vibration absorption peak of phosphoric acid-modified biochar (PBC) at 3400 cm^− 1 increases (Fig. 2), indicating that acid modification introduces more hydroxyl groups or makes the existing hydroxyl groups more exposed³⁴. Correspondingly, the intensity of the C-H stretching vibration absorption peak of PBC at 2900 cm^− 1 weakens, suggesting that acid modification removes some alkyl structures and reduces hydrophobic moieties. All the mentioned phenomena reveal that the hydrophilicity of the acid-modified biochar has been enhanced. The absorption peak of the C = O/C = C stretching vibration of PBC at around 1600 cm^− 1 is enhanced, indicating the introduction of more carbonyl or carbon-carbon double bonds. The probing functional groups are able to provide more active sites useful for the adsorption of certain substances containing polar groups. For instance, the oxygen atom in the carbonyl group can form hydrogen bonds or electrostatic interactions with molecules containing lone pairs of electrons, thereby enhancing the adsorption capacity^35,36. The increase in the intensity of the absorption peak of C-O stretching vibration at 1000–1300 cm^− 1 indicates that acid modification introduces more functional groups containing C-O bonds, such as ester groups and ether bonds³⁷. The existence of these functional groups will have a positive impact on the adsorption of polar substances^38,39. Furthermore, this indicates the enhancement of the adsorption capacity of acid-modified biochar for nutrient ions.

Fig. 2.

The FTIR spectra for biochar from spent mushroom substrates before and after acid modification in full range (4000-400 cm-1). BC, biochar; PBC, phosphoric acid-modified biochar.

Comprehensive analysis of BET surface area and microporous structure

The obtained results reveal that the pore structure of biochar has undergone systematic reconstruction after different treatments. Compared with CK without biochar addition, all biochar treatments exhibited a more complex pore distribution and a higher surface area. The results of pore structure analysis indicated that, compared with CK, the total porosity was significantly improved in all biochar treatments (Fig. 3). Compared with BMS, the volume of micropores and BJH pores under AMMS treatment increased significantly, and the average pore diameter decreased significantly. The surface area of the mushroom substrate after adding biochar increased exponentially (Table 2). The surface areas treated with B3 and PB3 were 29 times and 33 times that of CK, respectively. Compared with BMS, the average surface area of each treatment in AMMS increased significantly. Consistent with the previous research results on the effect of temperature and pretreatment processes on the physical properties of biochar^40,41acid modification significantly increased the micropore volume and BJH pore volume of the treated biochar, thereby also enhancing the specific surface area of AMMS. For mycelial growth, a larger surface area and more pores of the mushroom substrate mean a higher chance of spore attachment and better ventilation performance⁴². This indicates that AMMS has a good microstructure, which is of great significance for increasing the quality and yield of oyster mushroom^43,44.

Fig. 3.

The pore structural parameters of BMS and AMMS.

The characteristics of mushroom substrate during the growth period of oyster mushroom

The optimal substrate conditions for oyster mushroom growth include a moisture level of 60–65% and a pH range of 6.5–7.0. Deviations beyond these parameters can impede mycelium growth and hinder fruit body formation⁴⁵. In terms of pH, with the increase of the biochar additive, the pH of the mushroom substrate decreased significantly (Table 3). However, with the development of the fruiting bodies, the pH in each treatment substrate increased significantly. There is no significant pH difference between each treatment in AMMS and BMS. As agricultural residues decompose during mushroom mycelial growth, the production of organic acids occurs, resulting in a progressive decline in pH levels in the surrounding environment⁴⁶. However, the active surface moieties of the biochar play a crucial role in buffering the acidification process and ensuring pH stabilization⁴⁷.

The moisture of CK varies greatly. The moisture exceeded 70% at the first harvest but dropped sharply to around 50% at the second harvest. Correspondingly, the moisture variation ranges of each treatment in AMMS and BMS with biochar added were relatively small, both ranging from 60 to 65%. This indicates that biochar addition can improve the water holding and aeration capacity of oyster mushroom substrate, which is ascribed to the complex pore structure of biochar³. The EC of the mushroom substrate was significantly enhanced by adding biochar. Meanwhile, the EC of each treatment in AMMS was significantly higher than that of each treatment in BMS, with an average increase of about 30%. This means that PBC increases the availability of total water-soluble nutrient substances in the mushroom substrate⁴⁸. In conclusion, the mushroom substrate with biochar additive has more stable properties and improved water retention and air permeability. In addition, PBC further enhances the bioavailability of nutrients in the mushroom substrate and boosts the growth of its fruiting bodies.

Mycelial growth and oyster mushroom yield

In our previous work, we discovered that the addition of some amounts of biochar to the mushroom substrate can promote mycelial growth, enhance the yield of oyster mushrooms and brief the cultivation period²². According to Fig. 4A, the mycelium growth rate of the mushroom substrate with biochar added was dramatically more than that of CK. Moreover, the entire mycelium bag could be filled 16 days after spawning. Compared with the treatments of BMS, the treatments of AMMS can fill the mycelium bags for nearly 12–13 days, which indicates that AMMS can further promote the growth of mycelium. It is worth noting that the mycelia of both B3 and PB3 treatments were not covered with bacterial bags, indicating that the mycelial growth of these two treatments was inhibited. The growth status of mycelium can directly reflect the yield of oyster mushrooms. The yield of PB1 and PB2 treatments with the best mycelium growth was the highest among all mushroom substrates, on average 13–16% higher than that of B1 and B2 treatments, which ranked second (Fig. 4B). Meanwhile, the first mushroom harvest time of PB1 and PB2 was also the earliest, around 25 days, which was an average of 2.5 days earlier than that of B1 and B2 treatments. This indicates that acid-modified biochar has a stronger effect than unmodified biochar on promoting the mycelial growth and increasing the yield of oyster mushrooms (Fig. 5).

Fig. 4.

Yield and mycelial growth of oyster mushrooms with various treatments. (A) Mycelium growth in diverse mushroom substrates. (B) Yield (column) and fruiting time (line) of oyster mushrooms grown on diverse mushroom substrates. Different letters indicate a significant difference (P < 0.05).  * At the initial harvest, close to half of the mushroom baglogs in both the B3 and PB3 treatments exhibited no mushroom production. Additionally, no fruit was observed in these treatments during the subsequent harvest.

Fig. 5.

Comparison of the growth of oyster mushrooms in BMS and AMMS 25 days after spawn.

By acid modification, AMMS provides a growth environment that is more appropriate for the development of fruiting bodies due to the increase of the specific surface area⁴⁹pore structure and the number of surface oxygen-containing functional groups of biochar⁵⁰ as a result of acid modification. Besides, acid treatment also removes the ash or impurities clogged in the pores of biochar, thereby further exposing the pores, enhancing the specific surface area, and providing more space and attachment points for mycelial growth⁵¹. Among all the treatments, the yield and cultivation period performance of CK were the worst except for the B3 and PB3 treatments. As for these two treatments, the excessive addition of biochar led to overdosing the C/N ratio of the substrate to a level above the optimum quantity for oyster mushrooms, which inhibited thei growth⁵². Karlsson et al. demonstrated that incorporating two distinct levels of biochar into the cultivation substrate of oyster mushrooms negatively affected the yield of fruiting bodies⁵³. The elemental composition of the fruiting bodies mirrors that of the biochar, indicating that elemental enrichment could be one of the underlying reasons. This result aligns with the inhibitory effects observed in the B3 and PB3 treatment groups in this study, collectively underscoring the critical importance of optimizing the biochar addition concentration for practical applications.

In addition to yield and cultivation time, economic benefits are also an important factor that edible fungus producers pay attention to. In our previous research, the total benefit per ton of BMS was $221.5 higher than that of the common processing method³. In this study, through input-output analysis, the total benefit of AMMS was about 12% higher than that of BMS, which indicates that it is feasible in practice to recover spent mushroom substrate in the form of acid-modified biochar.

Mapping of heavy metal pollutants and metabolites within the mushroom substrate

The safety of adding biochar to cultivation substrates is an important issue that has attracted much attention in agricultural production⁵⁴. To investigate if the mushroom substrate composed of acid-modified biochar poses a food safety risk to oyster mushrooms, the pollutants and metabolites of the fruiting bodies were tested for possible risks. The results indicated that the NMR spectra of the samples treated with biochar addition were basically similar to those of CK, suggesting high similarity in the metabolome of all mushroom samples (Fig. 6). Meanwhile, the contents of pollutants in each treated sub-entity were all within the safety range stipulated by the national standards⁵⁵ (Fig. 7). This indicates that while AMMS has increased the yield of oyster mushrooms, its safety has also been verified.

Fig. 7.

NMR spectrum of oyster mushrooms cultured on BMS and AMMS.

Fig. 6.

Analysis of contaminant contents in oyster mushrooms cultured on BMS and AMMS. The dotted lines indicate the thresholds for pollutants, as stipulated in the "Food Safety National Standard-Limit of Contaminants in Food" established by the National Health Commission of the People’s Republic of China.

The use of biochar is not without risks. Depending on its source material, biochar has the potential to contain harmful pollutants, like soluble organics, heavy metals, and persistent free radicals⁵⁶which, if introduced into a product, can pose substantial health hazards to humans. Some biochar composed of raw materials even contains highly toxic metal cyanides, which can reach 85,870 mg/kg under extreme conditions⁵⁷. This study verified through experiments that neither BMS nor AMMS brings food safety risks to the production of oyster mushrooms.

Conclusions

Herein, we explored the influence of AMMS on the cultivation of oyster mushrooms. The obtained results reveal the role of acid modification in the significant enhancement of the specific surface area, pore structure and the number of oxygen-containing surface moieties of biochar, which further optimizes the physicochemical properties of the medium, and makes it more conducive to the mycelium growth of oyster mushrooms. The pH value and humidity of mushroom substrate containing PBS are more stable, the electrical conductivity is higher, and the bioavailability of nutrients is enhanced. The mycelium growth rate accelerated, the fruiting time advanced, the yield increased by 13%-16% compared with BMS, and the economic benefits improved significantly. The contents of heavy metal pollutants and metabolites in each treatment were all within the national safety standards and did not pose any food safety risks. In conclusion, AMMS is a new way for the efficient resource utilization of SMS. It also offers scientific basis and technical support for optimizing the cultivation techniques of oyster mushrooms, enhancing production efficiency and ensuring product safety.

Table 1.

Proximate and elemental content in AMMS.

Characteristics	CK	B1	B2	B3	PB1	PB2	PB3
Proximate content (wt%)
Volatile matter	57	34	22	14	29	21	16
Fixed carbon*	21	53	68	78	58	70	76
Ash	8	6	5	4	6	5	5
Water	14	7	5	4	7	4	3
Elemental content (wt%)
C	36	47	52	70	43	47	64
N	5	3	2	2	3	3	1
H	6	5	4	3	3	4	3
O**	53	45	42	25	51	46	32
S	0	0	0	0	0	0	0
O/C	1.47	0.95	0.8	0.35	1.18	0.97	0.5

* Calculated by sample weight loss (100 wt%-volatile matter-ash-water).

** Calculated by sample weight loss (100 wt%-carbon-nitrogen-hydrogen-sulfur).

Table 2.

The BET (Brunauer, emmett, and Teller) surface area of mushroom cultivation substrates of different treatments in BMS and AMMS. Different lowercase letters denote significant differences (P < 0.05).

Treatments	BET specific surface area(m2 g− 1)
CK	5.78 ± 0.26d
B1	32.71 ± 1.73c
B2	70.38 ± 3.41b
B3	169.52 ± 8.33a
PB1	40.46 ± 2.04c
PB2	81.46 ± 3.57b
PB3	189.72 ± 9.18a

Table 3.

Dynamic changes of mushroom substrate parameters during mycelial and fruiting body growth. All data were gathered prior to water stimulation.

Characteristics	Time	CK	B1	B2	B3	PB1	PB2	PB3
pH	4d	6.77 ± 0.12	6.60 ± 0.11	6.78 ± 0.12	6.94 ± 0.12	6.77 ± 0.13	6.58 ± 0.12	7.02 ± 0.15
	8d	6.51 ± 0.11	6.77 ± 0.13	6.45 ± 0.12	6.84 ± 0.11	6.65 ± 0.12	6.67 ± 0.11	6.76 ± 0.13
	12d	6.37 ± 0.11	6.68 ± 0.11	6.54 ± 0.11	6.67 ± 0.10	6.41 ± 0.11	6.80 ± 0.13	6.83 ± 0.12
	16d	6.00 ± 0.10	6.35 ± 0.09	6.64 ± 0.11	6.94 ± 0.14	6.30 ± 0.11	6.66 ± 0.12	6.68 ± 0.12
	20d**	6.19 ± 0.10	6.46 ± 0.11	6.45 ± 0.12	6.58 ± 0.12	6.22 ± 0.12	6.32 ± 0.12	6.91 ± 0.13
	2nd harvest*	4.85 ± 0.08	6.38 ± 0.12	6.52 ± 0.13	6.44 ± 0.11	6.19 ± 0.10	6.57 ± 0.11	6.50 ± 0.11
Moisture (%)	4d	60.31 ± 1.21	61.39 ± 1.23	60.43 ± 1.21	60.66 ± 1.21	61.50 ± 1.21	60.70 ± 1.21	61.50 ± 1.23
	8d	63.96 ± 1.34	61.91 ± 1.24	61.93 ± 1.24	65.45 ± 1.31	62.77 ± 1.26	64.30 ± 1.29	65.24 ± 1.30
	12d	66.92 ± 1.31	65.65 ± 1.31	62.47 ± 1.25	63.67 ± 1.27	65.82 ± 1.32	64.35 ± 1.29	64.84 ± 1.29
	16d	68.69 ± 1.37	65.42 ± 1.32	64.19 ± 1.28	61.09 ± 1.22	62.41 ± 1.23	61.58 ± 1.23	60.83 ± 1.22
	20d**	72.22 ± 1.44	65.10 ± 1.30	60.85 ± 1.22	61.34 ± 1.23	64.12 ± 1.28	62.50 ± 1.25	60.98 ± 1.22
	2nd harvest*	51.73 ± 1.44	58.61 ± 1.17	59.92 ± 1.20	60.87 ± 1.22	63.51 ± 1.17	62.28 ± 1.15	61.44 ± 1.19
EC (dS/m)	4d	10.12 ± 0.51	15.32 ± 0.77	15.95 ± 0.80	16.82 ± 0.84	21.33 ± 1.07	21.43 ± 1.07	21.72 ± 1.09
	8d	10.54 ± 0.53	15.44 ± 0.78	16.02 ± 0.82	17.12 ± 0.86	21.75 ± 1.09	21.56 ± 1.08	21.93 ± 1.11
	12d	10.78 ± 0.54	15.89 ± 0.79	16.23 ± 0.81	17.33 ± 0.87	21.82 ± 1.11	21.89 ± 1.09	22.01 ± 1.08
	16d	11.21 ± 0.56	16.12 ± 0.81	16.92 ± 0.85	17.54 ± 0.88	21.96 ± 1.10	22.06 ± 1.10	22.18 ± 1.12
	20d**	12.31 ± 0.62	16.83 ± 0.84	17.81 ± 0.89	17.89 ± 0.89	22.15 ± 1.07	22.37 ± 1.12	22.41 ± 1.12
	2nd harvest*	9.75 ± 0.49	14.61 ± 0.73	15.13 ± 0.76	16.57 ± 0.83	20.97 ± 1.05	21.29 ± 1.06	21.63 ± 1.08

* Owing to the initial inconsistency in the first harvest timing for each treatment, the subsequent harvest times differed as well, resulting in the final measurements being dependent on the actual harvest time for each treatment.

** After twenty days post-spawning, primordia emerged in certain treatments, requiring water for inducing regular fruiting, while others had to postpone. Consequently, the timing of water stimulation varied among treatments after the initial twenty days, rendering the collection of pH and moisture data unnecessary.

Author contributions

Wei Hu: Data curation, Writing–original draft, Lanming Gou: Methodology, Writing – review & editing, Liujie Hu: Investigation, Tao Liang: Methodology, Shuai Wang: Funding acquisition, Na Zhou: Conceptualization, Supervision.

Data availability

Raw data supporting the conclusions of this article will be made available from the corresponding author upon reasonable request.

Declarations

Competing interests

The authors declare no competing interests.