1. Introduction

Most plants form symbioses with mycorrhizal fungi since their emergence on land hundreds of millions of years ago (Brundrett, 2002; Genre et al., 2020; Martin and van der Heijden, 2024). Depending on carbon (C) and energy availability, these symbioses employ two nutrient acquisition strategies: (ⅰ) via mycorrhizal fungi and (ⅱ) via roots (hereafter as mycorrhizal and root strategies, respectively) (Brundrett, 2002; Chen et al., 2022; Han et al., 2024; Martin et al., 2017; Rich et al., 2021; Tedersoo et al., 2020; Zhang et al., 2023; Wang et al., 2025). The development and maintenance of root mycorrhizal symbioses is energy costly, particularly for trees that form ectomycorrhizas (ECM), with ECM mantle and emanating extramatrical hyphae consuming up to 22% and even more of plant net primary production (Drigo et al., 2010; Gamper et al., 2005; Han et al., 2021; Hobbie, 2006). Regardless of their evolutionary history, plants frequently experience C limitations, (e.g., as a consequence of a decline in atmospheric CO₂ concentration since the Cretaceous period) or current natural disturbances (e.g., leaf damage by insects, pathogens, or fire) (Barker et al., 2022; Comas et al., 2012; Fan et al., 2022; Fernandez et al., 2023). Consequently, studies on how and why plants shift between root and mycorrhizal strategies under C-limiting conditions are important for gaining an understanding of plant growth, evolution, interactions with fungi, and responses to environmental change.

Nutrient acquisition by mycorrhizas depends not only on the number of roots that form mycorrhizal associations (e.g., mycorrhizal colonization rate) but also on the composition of the fungal community (van Galen et al., 2023). For example, ECM communities dominated by fungi with lower C costs (e.g., shorter-distance hyphal exploration type or a thinner ECM mantle) can have higher nutrient benefits than ECM communities with higher C costs (e.g., longer-distance hyphal exploration type or a thicker ECM mantle) (Ding et al., 2023; Raven et al., 2018; Wang et al., 2022). Meanwhile, network properties of the fungal community may also be related to microbial function and tolerance to environmental stresses (de Vries et al., 2018; Hernandez et al., 2021; Hou et al., 2021; Wang et al., 2023; Zhai et al., 2024). For example, recent studies have shown that a more complex network can contribute to higher stability under conditions of environmental change (Liu et al., 2022; Yuan et al., 2021). Therefore, in addition to the strength of fungal associations, the community composition and network properties should be taken into consideration for a better understanding of fungal response and resistance to environmental stress, such as C limitation.

The root strategy for nutrient acquisition relies on both quantity (e.g., root surface area) and activity of absorptive roots (Wang et al., 2024; Zhang et al., 2024). The quantity of roots is determined by root biomass and length per unit root biomass [e.g., specific root length (SRL)], given that the root surface area is proportional to the length of the absorptive roots (Pregitzer et al., 2002). Activity-related root traits, such as root nitrogen (N) content and tissue density (Kramer-Walter et al., 2016), are widely reported independent of the quantity-related trait SRL (Bergmann et al., 2020; Carmona et al., 2021). Consequently, both quantity-and activity-related root traits should be taken into account when determining nutrient acquisition via the root strategy. However, few studies to date have concomitantly examined responses of the root and mycorrhizal strategies to C limitation by fully accounting for the key components of the two strategies; that is, the colonization, composition, and network of the fungal community, as well as root quantity and activity (e.g., root N content, respiration rate, and exudation rate).

In this study, we examined responses of roots and mycorrhiza to C limitation in mature ECM trees, Pinus taeda. Tree girdling was used to simulate the C limitation because it can cut off stem phloem by which photosynthate from leaves are transported belowground. We acknowledge some disadvantages of tree girdling, e.g., causing hurt to or even death of the plants besides introducing carbon limitation. However, tree girdling, unlike other methods for carbon limitation, e.g., shade or defoliation, has no effect on physical disturbance of the soil and the microorganisms therein (Hogberg et al., 2001), and the girdling also has no alteration on soil water, nutrients, and temperature (Table S1). It is well-known that tree girdling can also lead to nutrient limitation because nutrient acquisition by roots is energy-demanding (Beerling and Franks, 2010; Bennett and Groten, 2022), as such less nutrient absorption by roots under reduced photosynthate allocation belowground (Clausing et al., 2021; Pena et al., 2010). Nevertheless, this nutrient limitation as aforementioned is the indirect effect of carbon limitation. More importantly, such nutrient limitation not the commonly observed nutrient limitation that is usually caused by edaphic properties, e.g., soil parent material, soil erosion by wind or rainfall, etc. Therefore, we concentrate mainly on the effect of girdling-induced carbon limitation on nutrient acquisition strategies by roots and mycorrhizal fungi. Here, we hypothesized that tree girdling will cause a shift of nutrient acquisition from mycorrhizas to roots, given the high C costs entailed in maintaining mycorrhizal associations. To test this hypothesis, we analyzed a broad range of parameters associated with the root and mycorrhizal strategies and performed root transcriptome analysis to investigate molecular evidence for the predicted shift in nutrient acquisition strategies in response to the C limitation.

2. Materials and methods 2.1. Site description

This study was conducted at Madao Forestry Station, Biyang, Henan Province, China (113°42′E, 32°34′N; approximately 300 m a.s.l.) where Pinus taeda, ~25 years old, grew in monoculture, with sparse understory shrub species, Vitex negundo L. var. heterophylla (Franch.) Rehd. Mean annual temperature and precipitation are 14.6 ℃ and 960 mm, respectively. The soil belongs to Alfisol with soil pH 4.1, soil total C content 2.1% and soil total N content 0.18%.

2.2. Experimental design and sampling

On March 28, 2021, we selected mature Pinus taeda of similar size (average diameter at breast height = 22 cm) in a forest stand with relative homogeneous soil conditions, where each individual is spaced at a distance of 6–8 m. Five individuals were randomly chosen as the non-girdled trees, and the girdling was performed on another five individuals each with a distance over 20 m from the girdled ones. The full girdling of tree stems invariably causes the death of trees (Doughty et al., 2020), thus to avoid this outcome, each of the five trees was girdled by removing half of the bark in a 5 cm long section around the circumference of the stem at a height of 1.5 m above the ground. We further showed that such girdling had no effect on soil temperature, soil water and nutrient contents (soil total carbon and total nitrogen content) (Table S1).

In underground ecosystems, plant roots typically grow in an entangled manner, making it difficult to separate the absorptive roots of the same tree from those of other trees. Thus, we used the ingrowth root bag method to harvest absorptive roots and extramatrical hyphae from target roots. This frequently used method considerably reduces interference from the surrounding plant roots to the target roots and ensures a valid comparison with newly grown roots for the girdled and non-girdled trees (Eissenstat et al., 2015; Lekberg et al., 2013; Phillips et al., 2014). To examine the maximum effects of girdling on the roots and mycorrhizas, ingrowth root bags were placed on the girdled side of each tree. This is because allocation of current photosynthates to lateral roots growing in the girdled side is greatly reduced.

Briefly, we initially removed leaf litter from the trunk to expose the main woody roots of the targeted trees (surface soil to a depth of 10 cm). Woody lateral roots, ca. 4 mm in diameter and ca. 20 cm in length, were identified when tracking the main roots. One lateral root was pruned prior to being inserted into a nylon bag (20 × 30 cm; mesh size, 0.5 mm) filled with 2 kg of in situ sieved surface soil that was free of any visible plant materials. In addition, two small nylon bags (3 × 10 cm; mesh size, 48 μm) filled with 21 g of acid-washed quartz sand were placed into each ingrowth root bag to collect external hyphae. ECM fungal hyphae can grow into this medium, while saprophytic fungal hyphae cannot (Korkama et al., 2007; Lindahl et al., 2007; Wallander et al., 2010). These hyphae were mainly derived from the regrown lateral roots. After sealing the ingrowth root bags with a silicone sealant (Phillips et al., 2014), the bags were buried in the surface soil and covered with the leaf litter in situ. To ensure a sufficient quantity of roots and extramatrical hyphae, five root bags were placed at the girdled side of each tree individual.

Given that conifer roots regrew slowly, the root and hyphal samples were harvested after seven months of growth, by mid-October 2021. Unfortunately, no regrown lateral roots in each of the five bags were observed for one non-girdled tree individual, whereas for each of the other nine tree individuals, new lateral roots were observed in at least four of the five bags. Four ingrowth root bags with regrown roots for each tree individual were used for the following measurements, respectively: (1) root sample respiration, root carbon (C) and nitrogen (N) contents, mycorrhizal colonization, extramatrical hyphal length density and absorptive root biomass; (2) root exudates; (3) root anatomy and root non-structural carbohydrate; and (4) root transcriptome and composition of the root-colonizing fungal community (See Fig. 1).

The root samples used for anatomical measurements were immediately placed in a formalin-aceto-alcohol solution (90 mL of 50% ethanol, 5 mL of 100% glacial acetic acid, and 5 mL of 37% methanol). Meanwhile, the samples used for root transcriptome and fungal composition analyses were immediately frozen in liquid nitrogen and stored at −80 ℃ for later use. All measurements were conducted for the absorptive root segment; that is, the terminal 2−3 orders of a root branch, which play major roles in the absorptive function of roots (Guo et al., 2008).

2.3. Soil characteristics

Soil temperature and water content (v/v%) in the top 10 cm soil were measured using temperature and moisture sensors (Takeme-10EC; China). Soil total C and total N content were also measured using an EuroEA 3000 elemental analyzer (HEKAtech Gmbh, Wegberg, Germany).

2.4. Root sample respiration

Under field conditions, it is difficult to distinguish roots from mycorrhizal fungi. Therefore, the measurement of respiration of our root samples, i.e., root sample respiration, included the roots and associated mycorrhizal fungi. We defined the respiration by roots themselves as "root respiration", and the respiration by roots with mycorrhizal association as "mycorrhizal root respiration". Root sample respiration was measured from 11:00 to 14:00 on October 15th using a closed static chamber system and infrared gas analyzer (GMP343; Vaisala, Vantaa, Finland) (Makita et al., 2009; Sun et al., 2017a, 2021). Several intact and representative absorptive root segments, typically comprising a mixture of mycorrhizal and non-mycorrhizal root tips, were carefully cleaned to remove adhering soil and impurities. The roots were then washed twice–first with distilled water, and finally with sterilized water–to prevent damage. The cleaned roots were immediately placed into the chamber of the Vaisala CO₂ gas-measuring probe, which was connected to an NR-1000 data recorder (Keyence, Japan) to record the CO₂ concentration in the chamber at 1s intervals for approximately 15 min. To obtain a stable CO₂ efflux from the roots, we recorded the CO₂ concentration approximately 5 min after the roots placed into the chamber (Sun et al., 2021). The temperature within the chamber (17–21 ℃) was recorded simultaneously using a UT331 thermometer (Unitech Company, Ltd., China). Root sample respiration rate per unit root dry mass was calculated using the following formula:

where C_co2 is the CO₂ concentration (ppm) at a given time, n is the number of time intervals during the measurement period, t_i is the start time, Δt is the time interval, V_s is the volume of the chamber (0.144 l), 22.4 is the standard gas volume, T is the chamber temperature (℃), and W is the dry mass of the root sample (g). We also calculated the root sample respiration rate per unit root length using a similar formula. For a comparison of root sample respiration between the girdled and non-girdled trees, we adjusted the measured respiration rate to a respiration rate at 20 ℃ according to Tjoelker et al. (2001) to consider the effects of different chamber temperatures on the comparison of root sample respiration between treatments (Sun et al., 2021; Tjoelker et al., 2001). During the period for root respiration measurement, soil temperature and water content (v/v%) were measured using temperature and moisture sensors (Takeme-10EC; China) (Table S1).

2.5. Root morphology and biomass

After the measurement of root sample respiration, the roots were scanned, and photographs were analyzed using WinRHIZO (Regent Instruments, Quebec, Canada) to obtain values for total root length, root volume, and average root diameter. We then calculated the specific root length (SRL; root length per unit root dry mass) and root tissue density (RTD; root dry mass per unit volume). Root biomass in each ingrowth root bag was determined by drying samples (60 ℃, 24 h).

2.6. Root chemistry

After measuring root morphology, the root samples were then ground and sieved to determine root C and N contents using an EuroEA 3000 elemental analyzer (HEKAtech Gmbh, Wegberg, Germany). Non-structural carbohydrate including soluble sugars and starch, was determined using a modified phenol-sulfuric acid method (Fan and Guo, 2010; Guo et al., 2004).

2.7. Root carbon exudation

Root exudates were collected from 07:00 on October 15th to 07:00 on October 16th using a non-soil syringe system modified from that described by Phillips et al. (2008). This method is suitable for the field collection of root exudates from mature woody plants. Briefly, we initially selected intact absorptive root segments and carefully washed the roots using deionized water prior to placing these in a 50-mL syringe containing 15 g of acid-washed glass beads and 25 ml of C-free nutrient solution (0.1 mM KH₂PO₄, 0.2 mM K₂SO₄, 0.2 mM MgSO₄, and 0.3 mM CaCl₂) wrapped with aluminum foil (Phillips et al., 2008). After incubating the roots in the solution for 24 h, the solution was collected from the syringe. To ensure the complete recovery of exudate C, the syringe was rinsed twice using 10 mL of the aforementioned C-free nutrient solution, retaining the washing thus obtained. The recovered solution was then filtered through a 0.22-μm sterile syringe filter (Sartorius, Minisart, Göttingen, Germany) (Sun et al., 2017a, 2017b; Tanikawa et al., 2018). The total organic C content in this solution was determined using a total organic C analyzer (TOC-L CSN; Shimadzu, Japan) (Stubbins et al., 2015), and the root exudation rate was calculated by dividing the total organic C content by the incubation time and root dry weight (and root total length) in the syringe.

2.8. Mycorrhizal colonization

For each tree individual, 50 root tips from the girdling side were randomly selected to determine whether they were infected with ectomycorrhizal fungi using a Nikon ECLIPSE NI–U microscope (Nikon, Japan) at × 100 magnification. Root tips characterized by a yellowish brown to golden yellow color and swollen shape were considered mycorrhizal roots. Colonization rate was measured as the percentage of ectomycorrhizal roots in the total number root tips examined (Danielsen et al., 2013; Teste et al., 2014).

2.9. Extramatrical hyphal length density

Membrane filtration technology (Rillig et al., 1999) was used to extract extramatrical hyphae. Briefly, quartz sand (4.0 g) in a small nylon bag was placed in a solution of 100 mL deionized water and 12 mL sodium hexametaphosphate (35 g L⁻¹). After shaking by hand for 30 s and allowing it to settle for 30 min, the supernatant containing the hyphae was filtered through a 38-μm sieve. The hyphae collected on the filter were transferred to a flask containing deionized water, and then re-transferred from the flask onto a 1.2-μm filter (Kejin, China) for staining and observation under a Nikon ECLIPSE NI–U microscope (× 200 magnification) (Mc et al., 1990; Tennant, 1975). Hyphal length density (HLD) was calculated using the grid intersection method according to the following formula:

where 0.03928 is the coefficient of grid conversion growth in cm, N is the number of cross-points between the mycelium and grid, 100 is the fractionation multiple (2 ml of 200 ml filtrate), and A is the field of view area (mm²) observed on the filter membrane (Tennant, 1975).

2.10. Ectomycorrhizal mental thickness

Ectomycorrhizal mantle thickness was determined after measuring the root anatomical structures following the protocols described by Guo et al. (2008) and Kong et al. (2014). Briefly, 15 root tips were randomly selected for each tree, embedded in paraffin, cut into 8-μm thick sections for staining, and then photographed using a Nikon ECLIPSE NI–U microscope. Mantle thickness and its contribution of mantle thickness to root diameter (Mantle %) (Withington et al., 2006) were determined using ImageJ software (NIH Image, Bethesda, MD, USA).

2.11. Fungal composition and network in roots

We used high-throughput sequencing to determine the species composition of absorptive root fungi. Generally, this analysis involves DNA extraction, PCR amplification, and sequencing. Genomic DNA of the microbial community was extracted from the root samples using the E.Z.N.A.® soil DNA kit (Omega Bio-tek, Norcross, GA, U.S.) according to the manufacturer's instructions. DNA concentration and purity were determined using a NanoDrop 2000 UV–vis spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). Amplicon fragments were amplified using the primer pair ITS1F (5′-CTTGGTCATTTAGAGGAAGTAA-3′) (Gardes and Bruns, 2008) and ITS2R (5′-GCTGCGTTCTTCATCGATGC-3′) (White, 1990), targeting the fungal ITS1 region of ribosomal DNA. The PCR amplification procedure was as follows: initial denaturation at 95 ℃ for 3 min, followed by 35 cycles of denaturation at 95 ℃ for 30 s, annealing at 55 ℃ for 30 s, and extension at 72 ℃ for 45 s, and a single extension at 72 ℃ for 10 min, and then held at 10 ℃. Finally, purified amplicons were pooled in equimolar amounts and paired-end sequenced using the Illumina MiSeq PE300 platform (Illumina, San Diego, CA, USA) according to the standard protocols of Majorbio Bio-Pharm Technology Co. Ltd. (Shanghai, China). The raw sequencing data have been uploaded to the National Center for Biotechnology Information BioProject database under accession number PRJNA918806.

The raw gene sequencing reads were demultiplexed, quality-filtered using fastp version 0.20.0, and merged using FLASH v.1.2.7 (Chen et al., 2018) with the following criteria: (ⅰ) Filter reads with a tail mass value below 20 and set a 10bp window. If the average quality score of < 20, cut off the back base from the window. Filter reads with a quality control value below 50bp and remove reads containing N bases. (ⅱ) Only overlapping sequences longer than 10 bp were assembled according to their overlapped sequence. The maximum mismatch ratio of overlap region was 0.2. Reads that could not be assembled were discarded. (ⅲ) Samples were distinguished according to the barcode and primers, and the sequence direction was adjusted, exact barcode matching, 2 nucleotide mismatches in primer matching.

Operational taxonomic units (OTUs) with a 97% similarity cut-off were clustered using UPARSE version 7.1, and chimeric sequences were identified and removed (Edgar, 2013). The taxonomy of the fungal sequences was determined using The Ribosomal Database Project Classifier algorithm against the United States database (v.7.0/its-fungi database) using a confidence threshold of 70% (Wang et al., 2007). We used FUNGUILD to annotate the functional types of fungi, such as ECM, pathogen, and saprotroph (Nguyen et al., 2016). ECM fungi at the genus level were further classified into four mycorrhizal hyphal exploration types (contact hyphal exploration type, short-distance hyphal exploration type, medium-distance hyphal exploration type and long-distance hyphal exploration type) based on genus information according to the DEEMY database and references in Agerer (2001, 2006), Tedersoo and Smith (2013), and Guo et al. (2021).

For both the girdled and non-girdled trees, we also analyzed the complexity of the fungal network in the roots according to recent studies (Wu et al., 2022; Yuan et al., 2021). The complexity of networks can be evaluated based on network nodes, links, average degree, average weighted degree, and network density.

2.12. Root transcriptome sequencing

Total RNA was extracted from roots using TRIzol® reagent according to the manufacturer's instructions (Invitrogen, Carlsbad, CA, USA), and genomic DNA was removed using DNase I (TaKaRa, Japan). The integrity and purity of the total RNA were determined using a 2100 Bioanalyzer (Agilent Technologies, Inc., Santa Clara, CA, USA) and an ND-2000 NanoDrop spectrophotometer (Thermo Scientific). RNA purification, reverse transcription, library construction, and sequencing were performed according to the manufacturer's instructions (Illumina) at Shanghai Majorbio Bio-pharm Biotechnology Co., Ltd. Root RNA sequencing libraries were constructed using an Illumina TruSeqTM RNA sample preparation Kit and sequenced using the Illumina NovaSeq 6000 sequencing platform.

Raw reads were trimmed, quality control was performed using SeqPrep (https://github.com/jstjohn/SeqPrep) and Sickle (https://github.com/najoshi/sickle), and the clean data were used for de novo assembly using Trinity (http://trinitynaseq.sourceforge.net/) (Grabherr et al., 2011). All transcripts from transcriptome sequencing were annotated based on six commonly used databases [RefSeq non-redundant proteins (NR), Swiss-Prot, Pfam, Clusters of Orthologous Groups of proteins (COGs), Gene Ontology (GO), and Kyoto Encyclopedia of Genes and Genomes (KEGG)] for functional determination. To identify genes that were differentially expressed (DEGs) in response to girdling, we initially determined the expression levels of transcripts using RSEM software (http://deweylab.github.io/RSEM/). The DEGs between the girdled and non-girdled trees were determined using DESeq2 software (http://bioconductor.org/packages/stats/bioc/DEGSeq2/) with FDR (False discovery rate) ≤ 0.05, DEGs with |log2FC| > 1 and FDR ≤ 0.05 were considered to be significantly differential expressed genes. Finally, GO and KEGG were used to identify DEGs that were significantly enriched in GO terms and metabolic pathways, respectively, at a corrected p-value ≤ 0.05. The raw data have been uploaded to the National Center for Biotechnology Information BioProject database under accession number PRJNA918806.

2.13. Statistical analyses

Differences in the root and mycorrhizal traits between the girdled and non-girdled trees were analyzed using one-way ANOVAs (in conjunction with Tukey's HSD test). Pairwise root-mycorrhizal relationships were assessed using linear regressions. We also used principal component analysis (PCA) to assess the multiple variable relationships for root and mycorrhizal traits. Fungal community diversity for the girdled and non-girdled trees was compared using alpha_diversity.py in Quantitative Insights into Microbial Ecology (QIIME). Data were transformed to meet the requirements of normal distribution (e.g., root sample respiration rate was reciprocally transformed and root exudation rate was log-transformed) and equal variance of the data for the above analyses. These analyses were performed using R 4.1.0 (R core Team) with significant differences defined at p < 0.05, and a marginal significance at p < 0.1. The topological properties of the fungal network were analyzed using the psych and igraph packages in R software (R Core Team), and the network was illustrated using Gephi 0.9.2. The robustness and ZiPi (Zi: within-module connectivity; Pi: participation coefficient) of the fungal network were analyzed using the ggClusterNet package in R software (R Core Team). Differences in the root transcriptomes between the girdled and non-girdled trees were analyzed using the I-Sanger Shengxin Cloud Platform (Shanghai Meiji Biomedical Technology Co., Ltd.).

3. Results 3.1. Root strategy responses

There was no difference between the control and girdled trees with respect to absorptive root biomass or SRL (Fig. S1a, b). However, compared with the non-girdled trees, root sample respiration rate in girdled trees increased by 67% (Fig. 2a; Fig. S2a), whereas significant reductions were detected in root exudation rate (Fig. 2b; Fig. S2b), root N content (Fig. 2c), root C content (Fig. 2d), and root non-structural carbohydrate content (Fig. 2e) in response to the girdling. In addition, there was an increase in the ratio of root N content to root non-structural carbohydrate content following the girdling (Fig. 2f).

3.2. Mycorrhizal strategy responses

In response to girdling, mycorrhizal colonization rate, extramatrical hyphal length density and hyphal length density per unit root sample respiration increased by 106%, 342% (Fig. 2g and h), and 162% (Fig. 2i), respectively. However, ectomycorrhizal mantle thickness and the percentage of root diameter contributed by the mantle were unaffected by the girdling (mantle %, Fig. S3).

3.3. Relationships within and between root and mycorrhizal strategies

Across the nine tree individuals, root sample respiration rate increased with a reduction in exudation rate, whereas mycorrhizal colonization rate increased with extramatrical hyphal length density (Fig. 3a; Table S2). In addition, hyphal length density per unit root sample respiration was positively correlated with mycorrhizal colonization (r = 0.64, p = 0.064) (Fig. 3b). Across the root and mycorrhizal strategies, root sample respiration rate increased and root C content (41–52%) declined with an increase in mycorrhizal colonization and extramatrical hyphal length density (Fig. 3a–c, d; Table S2). Furthermore, absorptive root biomass was negatively correlated with mycorrhizal colonization (Table S2).

3.4. Responses of mycorrhizal composition

Girdling was found to be associated with a reduction in the number of fungal OTUs in roots (Fig. S4), although it had no significant effect on fungal diversity (Table S3). Girdling also promoted a reduction in the relative abundance of pathogenic fungi, although it had no apparent influence on the abundance of ectomycorrhizal fungi (Fig. 4a). Among the mycorrhizal fungi colonizing roots, we detected a marked increase in the relative abundance of the Contact-Medium hyphal exploration type (e.g., Russula sp. and Tomentella sp.), whereas a reduction was detected in the relative abundance of the Medium-Long hyphal exploration type (e.g., Amphinema sp.) (Fig. 4b; Table S4).

Further analysis using a volcano plot revealed that the three shorter-distance hyphal exploration types fungi, i.e., Russula sp., Sebacina sp., and Tomentella sp., were significantly upregulated after girdling (Fig. 4c). Among them, Russula sp. and Tomentella sp. are the Contact-Medium hyphal exploration type, while Sebacina sp. is the Contact-Short hyphal exploration type. In addition, a pathogen genus, Fusarium sp., was downregulated.

3.5. Fungal community network in roots

Girdling altered network properties (Fig. 5a and b; Fig. S5; Table S5) and the composition (Fig. 5c) of the fungal community. A suite of key trait values describing the network complexity of the fungal community in the roots was reduced after girdling (Fig. 5a), including the number of nodes and edges, average weighted degree, density, and number of positive and negative correlations (Fig. 5a). To determine the resistance of the microbial network to disturbances, a natural connectivity analysis was carried out. Natural connectivity of a complex network was applied to reveal the robustness of a network. We found that the natural connectivity values dramatically increased after the girdling (Fig. 5b), indicating weakened resistance. The ZiPi analysis at the genus level did not identify any key fungal genus that impact the overall network structure of the microbial communities (Fig. S6). Ectomycorrhizal fungi that were most sensitive to girdling in our volcano plot analysis (e.g., Russula sp., Sebacina sp., and Tomentella sp., see Fig. 4c) are not key nodes in the network.

3.6. Responses of the root transcriptome

Many key genes associated with root C metabolism were differentially expressed in response to girdling, including those involved in glycolysis (e.g., hexokinase, phosphofructokinase, and pyruvate kinase) and the tricarboxylic acid cycle (e.g., citrate synthase, malate dehydrogenase, and isocitrate dehydrogenase) for carbohydrate decomposition into CO₂ (Figs. 6 and 7; Fig. S7). In addition, lipid synthesis pathways in the plastid (e.g., fatty acid synthase) and the Kennedy pathway in the endoplasmic reticulum (e.g., long-chain acyl-coenzyme A synthetases, glycerol 3 phosphate acyltransferase, and phosphatidate phosphatase) in the roots of P. taeda (Figs. 6 and 7) were also significantly affected by the girdling.

4. Discussion 4.1. The shift from roots to mycorrhiza strategy following girdling

In contrast to our initial hypothesis, we observed a shift in nutrient acquisition strategy from roots to mycorrhiza in response to the belowground carbon (C) limitation induced by girdling. This response is supported by the marked increase in the mycorrhizal colonization rate and extramatrical hyphal density, as well as a reduction in root activity (Fig. 2; Fig. S2). For example, a reduction in root physiological activity after girdling is indicated by a concomitant reduction in the root N content and the rate of root exudation. These responses are assumed to be associated with a marked reduction in C allocation to roots, as evidenced by the lower non-structural carbohydrate content in roots (Fig. 2d and e) (Clausing et al., 2021), thereby providing insufficient substrates to fuel the energy-demanding processes of root nutrient acquisition (Clausing et al., 2021; Johnson and Edwards, 1979; Parker et al., 2017; Schneider et al., 2017).

Interestingly, we detected a substantial increase in root sample respiration rate, a key trait indicating root activity, in response to girdling (Fig. 2a; Fig. S2a). The root sample respiration we measured here includes the release of CO₂ from both the roots and mycorrhiza colonizing roots (Sun et al., 2017a; Trocha et al., 2010). Root sample respiration increased in response to the increase of mycorrhizal colonization and extramatrical hyphal length (Fig. 3a–d). Therefore, it is likely that the observed increase in root sample respiration subsequent to girdling arose from a higher consumption of the limited C via mycorrhizal root respiration (i.e., the respiration by roots with mycorrhizal association). The great C allocation to mycorrhizal association after the girdling may led to a reduced root respiration (i.e., respiration by roots themselves) rate as indicated by lower root N content and lower rate of root exudation (Fig. 2b and c; Fig. S2b). Collectively, the increase in belowground C allocation to mycorrhizal fungi underlies the preference for mycorrhizal rather than root strategy under the girdling-induced C-limited conditions.

Along with an increase in mycorrhizal colonization rate, girdling also had the effect of altering the composition of mycorrhizal fungi within the roots by promotion a shift toward a considerably higher proportion of the shorter-distance hyphal exploration types and a lower proportion of the longer-distance hyphal exploration types (Fig. 4). A shift in the mycorrhizal composition of this nature is considered to reflect an adaptation of the mycorrhizal strategy to C-limited conditions, based on a preferential selection of shorter-distance hyphal exploration types usually conferred with lower construction costs for per unit nutrient acquisition (Anthony et al., 2022; Chen et al., 2021; Guo et al., 2021; Saikkonen et al., 1999; Wasyliw et al., 2020; Xie et al., 2024). We further identified three ectomycorrhizal fungal genera, i.e., Russula sp., Sebacina sp., and Tomentella sp. (Fig. 4c), that belong to the shorter-distance hyphal exploration types (Table S4) and were significantly altered after the girdling, especially Russula sp. and Tomentella sp. with remarkably increased abundance (Fig. 4b). Moreover, given the higher metabolic activity of the shorter-distance exploration types, this preferential selection can also contribute to an increased root sample respiration after girdling, compared with that of the longer-distance hyphal exploration types (Agerer, 2001; Courty et al., 2010; Ding et al., 2023; Finlay, 2008).

4.2. Fungal community: C utilization and network complexity

Few studies have concomitantly examined the responses of roots and mycorrhizas to girdling-or defoliation-induced C limitation. Nevertheless, studies with similar periods after girdling or defoliation did observe a strong mycorrhizal association with roots (Binkley et al., 2006; Clausing and Polle, 2020; Cullings et al., 2008; Kuikka et al., 2003; Pena et al., 2010; Wasyliw et al., 2020). These observations are in line with our finding of increased dependence on mycorrhizal strategy under C limitation in P. taeda.

Given the high C cost of mycorrhizal associations, it would appear counterintuitive to shift to the mycorrhizal strategy when belowground C supply is reduced by girdling. Recent studies found that the association of roots with arbuscular mycorrhizal fungi (AMF) can reduce the C cost of nutrient acquisition by approximately 50% (Peng et al., 1993; Terrer et al., 2018; Ven et al., 2019). This suggests that C-use efficiency of nutrient acquisition (CENA, amount of nutrient acquisition per unit C cost) of AMF is higher than that of roots. Consequently, the associations of roots with AMF can conferred them with higher CENA, which could particularly be the case under C-limiting conditions (Raven et al., 2018; Wang et al., 2022). Similarly, the symbioses with ectomycorrhizal fungi (which typically occurs in low-fertility soils) may also promote an increase in CENA (Fig. 2f–i). For example, the C cost of ectomycorrhizal root sample respiration was twice that of non-ectomycorrhizal roots (Rygiewicz and Andersen, 1994), whereas nutrient acquisition by ectomycorrhizal roots can be 10-fold higher than that of non-mycorrhizal roots (Clausing and Polle, 2020).

Although there is no direct evidence for a higher CENA of ectomycorrhizal fungi than that of roots in P. taeda, we did detect a higher ratio of root N content (RN) to root non-structural carbohydrate content (NSC) after girdling (Fig. 2f) together with higher ectomycorrhizal colonization, which indirectly support this postulation. The rationale is that a higher CENA of ectomycorrhizal fungi than roots mean less C costs for per unit nutrient acquisition; this will cause relative less NSC consumption in per unit nutrient acquisition and hence more NSC left in roots, and hence a higher value of RN/NSC (Fig. 2f). Another evidence comes from the increased extramatrical hyphal length density (HLD)/root sample respiration after girdling (Fig. 2i). The higher value of HLD/root sample respiration means more hyphae production for nutrient acquisition per unit C cost via root sample respiration, that is, a higher CENA for the ectomycorrhizal fungi. Given the great difficulty in separating ectomycorrhizal fungi from roots in the field conditions, the higher CENA after girdling may come from (1) higher CENA of the ectomycorrhizal fungi themselves than the roots, and (2) a shift to the ectomycorrhizal communities of shorter-distance exploration types (e.g., Russula sp., Sebacina sp., and Tomentella sp., see Fig. 4c) usually with higher CENA (Kiers et al., 2011; Pena et al., 2010). Finally, the preference for mycorrhizal strategy in the girdled trees could represent a conservative strategy for nutrient cycling under C-limited conditions, in which nutrients derived from dying roots could be efficiently released and reused by symbiotic fungi with a high CENA.

In addition to the pronounced changes in fungal community composition in response to girdling, we also detected a reduction in the network complexity of the fungal community in the colonizing roots (Fig. 5a and b). A lower network complexity is taken to be indicative of greater instability and a lower resistance to environmental stress (de Vries et al., 2018; Pena et al., 2010; Yuan et al., 2021). Therefore, our result on the fungal network complexity suggests that increased dependence on mycorrhizal strategy for nutrient acquisition under C limitation conditions could come at a cost of reduced resistance to environmental stress.

Although we identified three ectomycorrhizal fungal genera (Russula sp., Sebacina sp., Tomentella sp.) potentially responsible for the change of CENA after the girdling, ZiPi analysis suggests that these fungi are not the key node microorganisms that impact the overall network complexity of the microbial communities (Fig. S6). Nevertheless, we could not rule out the critical role of these ectomycorrhizal fungi in determining the network complexity and stability given that ZiPi is a theoretical and mathematical analysis. Empirical studies could be employed to test the role of these fungi in affecting the microbiome network properties and root function.

4.3. Molecular mechanisms underlying the shift to mycorrhizas

Root transcriptome analyses can provide valuable insights into the molecular mechanisms underlying the shift in nutrient acquisition strategies between roots and mycorrhizas. For example, the downregulation of phosphoenolpyruvate carboxylase (PEPC) (Fig. 6) would tend to indicate a reduction in root activity in the tricarboxylic acid cycle, as lower PEPC levels could contribute to a reduction in the supply of substrates (oxaloacetic acid and malic acid) into the tricarboxylic acid cycle (Cao et al., 2021; Clausing et al., 2021; O'Leary, 1982), This indicates that the respiration by roots themselves is reduced, which is consistent with the reduced root physiological activity observed following girdling (Fig. 2b and c). In contrast, give the fatty acid auxotrophy of mycorrhizal fungi, the transfer of plant-derived fatty acids to fungi is necessary to sustain mycorrhizal associations (Bravo et al., 2017; Jiang et al., 2017; Luginbuehl et al., 2017). Our root transcriptome analysis in the present study revealed prominent changes in the metabolic pathways for lipid biosynthesis (Fig. 6). Consequently, the enhanced mycorrhizal association in response to girdling suggests an increased in lipid biosynthesis and/or transport to the ectomycorrhizas. Girdling may thus initiate a shift from the degradation of root carbohydrates via mitochondrial tricarboxylic acid cycling to lipid metabolism pathways (fatty acid biosynthesis in plastid and the Kennedy pathway for phospholipid biosynthesis in the endoplasmic reticulum) to sustain the increased dependence on mycorrhizas after girdling (Fig. 7).

5. Conclusions

By integrating multi-level evidence, including data obtained for root morphology, physiology, and transcriptome analyses, as well as the properties of the fungal communities colonizing roots, we demonstrated an unexpected shift in the nutrient acquisition strategy, from roots to mycorrhiza of a pine species under the girdling-induced carbon-limiting. Such a shift likely arises from the higher carbon (C) efficiency of nutrient acquisition (CENA) of mycorrhiza compared with that of roots, which hitherto has been greatly overlooked. Consequently, in response to girdling, belowground C allocation may shift towards lipid biosynthesis to sustain the increased dependence on mycorrhizal association. Interestingly, there could be a tradeoff between CENA and the stability of the fungal community colonizing roots; whereby roots subjected to C limitation harbor fungi with a higher CENA but the stability of the fungal community is reduced.

Overall, our study reveals an unrecognized key dimension for the roots–mycorrhiza interactions: CENA vs. the stability of the fungal communities in roots. This trade-off could be potentially be attributed to three ectomycorrhizal fungi belonging to shorter-distance hyphal exploration types, i.e., Russula sp., Sebacina sp. and Tomentella sp. This may pave a new way for future studies on the co-evolution between roots and mycorrhiza, the critical roles of mycorrhiza in adjusting soil organic C pool, and their responses to environmental changes. Practically, our findings will benefit plant restoration practices under stressful conditions, e.g., through inoculating roots with diverse microbiomes with a high CENA to increase network complexity and stability.

Acknowledgements

We thank Yawei Dong, Yu Tian, Yueshuang Hou, Haojie Wang for their help with the trait measurements in the field and lab. We also thank Prof. Yu Shi for his great help in microbial network analysis, Dr. Lijuan Sun for the revision of the early draft, and Mr. Shukui Zhao and Mr. Zhuo Guan for their support in the field work. We much appreciate two anonymous reviewers especially the handling editor for the insightful comments and suggestions in the microbiome analyses. This study was funded by the National Natural Science Foundation of China (32471824, 32171746, 42077450, 31870522, 31670550, 42122054), the leading talents of basic research in Henan Province, the Scientific Research Foundation of Henan Agricultural University (30500854), Excellent Youth Creative Research Group Project in Henan Province (252300421002), Foreign Scientists Studio in Henan province (GZS2025011), the Funding for Characteristic and Backbone Forestry Discipline Group of Henan Province, Guangdong Provincial Key Laboratory of Soil and Groundwater Pollution Control (2023B1212060002), Guangdong Basic and Applied Basic Research Foundation (2021B1515020082), the RUDN University Strategic Academic Leadership Program, Funding for Characteristic and backbone forestry discipline group of Henan Province and Research Funds for overseas returnee in Henan Province, China. We thank Editage Service in improving the English language.

CRediT authorship contribution statement

Jing Chen: Writing – review & editing, Writing – original draft, Software, Methodology, Investigation, Data curation. Jingjing Cao: Investigation. Binglin Guo: Methodology. Meixu Han: Methodology. Zhipei Feng: Methodology. Jinqi Tang: Methodology. Xiaohan Mo: Methodology. Junjian Wang: Writing – review & editing, Methodology. Qingpei Yang: Methodology. Yuxin Pei: Methodology. Yakov Kuzyakov: Writing – review & editing. Junxiang Ding: Writing – review & editing, Formal analysis. Naoki Makita: Writing – review & editing. Xitian Yang: Writing – review & editing. Haiyang Zhang: Writing – review & editing. Yong Zhao: Writing – review & editing. Deliang Kong: Writing – review & editing, Project administration, Funding acquisition, Formal analysis, Conceptualization.

Data availability

Data are available in the Dryad Digital Repository, a publicly available database, or can be acquired on request to the corresponding author.

Declaration of competing interest

We declare that we have no conflict interest.

Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.pld.2025.02.004.