Introduction

Global industrialization has led to an increase of tropospheric carbon dioxide (CO2) concentration from approximately 280 ppm in pre-industrial times to approximately 380 ppm nowadays, and it is expected to continue increasing in the future1,2. Alongside, the average surface ozone (O3) concentration has increased from an estimated pre-industrial value of 10 ppb to 20–45 ppb in the mid-latitudes of the northern hemisphere at a rate of 0.5–2% per year over the last decades2,3,4.

Industrialization has further led to a global pollution by organic pollutants, including polyaromatic hydrocarbons (PAHs)5. PAHs have mutagenic and carcinogenic properties, and show a high persistency in the environment6,7. Due to PAH contamination, huge areas are not suitable for agriculture or livestock anymore, and remediation of PAHs from soils is a priority goal to ensure food safety8,9. Among the proposed approaches, phytoremediation appears as an efficient and environment-friendly approach to remove PAHs from soils from large surfaces10. In most cases, phytoremediation of PAHs from soils was conducted by a single plant species2, PC1 microbial PLFAs), it led to a decrease in Gram-positive bacteria, a microbial group linked to PAH degradation49 and the most important microbial group involved in the removal of PAHs from soil in the present study (Fig. 2). Furthermore, eCO2 altered the soil microbial community composition, which is also in line with previous studies38,50,51 and calls for more detailed investigations of shifts in soil microbial communities with sequencing techniques.

We propose that this effect of elevated tropospheric CO2 may be due to the higher plant carbon input in soil resulting from enhanced photosynthesis52. This may lead to higher soil microbial activity29,53, as the pool of labile soil C may be increased by elevated root exudation33,54,55. In the present study, eCO2 had non-significant effects on the biomass of fungi and Gram-negative bacteria, but decreased the biomass of Gram-positive bacteria (Fig. S1). Consistent with the present study, both Larson et al.31 and Grueter et al.56 found that eCO2 had no significant influence on microbial biomass and activity, while Manninen et al.57 found a negative effect of eCO2 on soil microbial biomass. These variable results indicate that eCO2 effects on soil microbial communities may depend on the environmental context, such as soil conditions and/or plant community composition38.

Elevated O3 decreased plant biomass, soil microbial biomass and activity, and PAH removal. Ozone is a toxic compound that can induce oxidative stress in plants, and high tropospheric O3 concentrations have been reported to decrease inputs and to change the composition of assimilates into the rhizosphere34, which in turns affects soil microbial communities. Results of the present study indicate that ozone-mediated changes in soil communities may have dramatic effects on soil self-cleaning potential. Consistent with past studies58,59,60, a strong decrease in the biomass of Gram-positive and Gram-negative bacteria and shifts in microbial community composition in response to eO3 was observed (Figs 2, S1).

The effects of eCO2, eO3, and plant diversity on PAH removal were mediated to some extent by alterations of soil enzymatic activity. Both elevated CO2 and O3 led to a decrease in polyphenol oxidase activity, while plant functional group richness increased polyphenol oxidase but decreased phenol oxidase. In line with the present study, eCO2 reduced the activity61 and abundance62 of polyphenol oxidase, suppressed phenol oxidase63, while enzymes including phenol oxidase were strongly affected by plant species richness64. These results indicate that simultaneous alteration of plant community composition and environmental conditions may have contrasting effects on enzyme activity involved in PAH removal. Notably, many enzymes are involved in the metabolism process of PAHs65,66, some of which were not measured here. Although the measured enzymes responded significantly to the treatments, this did not explain variation in PAH removal, which is why they were not considered in the structural equation model (Fig. 2).

Elevated CO2 and O3 concentrations and variations in plant diversity had significant interactive effects on plant biomass, soil microbial functions, and the degradation of PAHs. Plant diversity altered the effect of eCO2 on soil microbial biomass and activity, but the clear positive interaction effects as expected in hypothesis (4) were not detected. This highlights the importance of plant diversity and community composition in mediating soil microbial functions in a future world, but also calls for a better mechanistic understanding of interactive effects of plant diversity and global change drivers.

However, plant diversity did not alter eCO2 and eO3 effects on PAH removal in the present study. This is in line with a recent meta-analysis by Thakur et al.46 showing no interactive effects of plant diversity and global change factors in affecting soil microbial biomass in the short term. Potentially, plant diversity-induced differences in soil microbial community composition and subsequent effects on essential services like PAH degradation need a longer time than captured by the present experiment46. Moreover, we propose that lum** plant community composition into functional group richness may not provide the adequate explanatory level. Instead, we propose that future studies may use more targeted plant trait-based approaches67, e.g., by considering root/rhizosphere traits, to develop a better mechanistic understanding of the relationship between plant community composition and functioning of soil communities linked to pollutant removal.

Importantly and in contrast to our hypothesis (4), eCO2 amplified the inhibitory effect of eO3 on PAH removal. This effect was partly mediated by an enhancement of eO3 effects on most soil microbial groups at elevated CO2. It particularly amplified the negative effect of eO3 on Gram-positive bacteria, the most important microbial group driving the removal of PAHs from soil in this study. This result exemplifies how different global change drivers can have unexpected synergistic effects on soil functions and compromise important ecosystem services.

Conclusion

We highlight that global environmental change factors, such as human-induced alterations in tropospheric gas composition, may undermine the ability of ecosystems to degrade pollutants. Soil self-cleaning showed a high robustness to alterations in plant diversity and community composition, yet elevated CO2 and O3 concentrations may compromise efforts such as phytoremediation to restore polluted soils. On the other hand, the present study also indicates that the targeted assembly of plant communities applying a more comprehensive knowledge regarding plant effects on soil biota may be a promising tool to shape soil microbial communities for the degradation of organic pollutants.

Materials and Methods

Open top chambers

The open top chamber (OTC) system is located at **anlin campus, Nan**g University, Nan**g, China (118°57′36.15″E, 31°7′23.99″N). Briefly, this system consists of four chambers with full control of atmospheric CO2 and O3 concentrations: one chamber with ambient CO2 (aCO2) and ambient O3 (aO3) levels, one with eCO2 and aO3 levels, one with aCO2 and eO3 levels, and one with both eCO2 and eO3 levels. The glass chambers are octagonal with 2 m in diameter and 2.8 m in height. CO2 was released from a tank (Q/JB-THB002, Bei**g Tianhai Industry Co., Ltd.), and O3 was produced by an O3 generator (NPF10/W, Shandong Lvbang Ozone Co., Ltd.) from pure O2. CO2 and/or O3 were mixed with air from temperature-controlled rooms and conveyed by fans (SFG-2, Shanghai Jiabao Co., Ltd.) to the bottom of the chambers. Gases were released into the antra via tiny holes in the stainless steel plate between the bottom and the antrum, and then released into the air of the open top of chambers. The quantity of the CO2 and O3 release was controlled by a flowmeter (LZB-3WB, Changzhou Shuangbo Co., Ltd.), the concentration of CO2 was detected with a CO2 monitor (Li-7000, Li-Cor, USA), and the concentration of O3 was detected with an O3 monitor (Model 205, 2B Co., USA). The O3 fumigation was conducted between 9:00 a.m. and 5:00 p.m. until harvest, except during rain events, and the CO2 fumigation was all day long until harvest. The target CO2 concentration for the eCO2 treatment was 200 ppm higher than aCO2, and the target O3 concentration for the eO3 treatment was 50–60 ppb higher than aO3 in order to simulate the forecasted tropospheric CO2 and O3 levels in 20502.

Plant cultivation

Three species of grasses (Lolium perenne, Dactylis glomerata, Phleum pratense), herbs (Plantago lanceolata, Taraxacum officinale, Centaurea jacea), and legumes (Trifolium pratense, Trifolium repens, Medicago sativa) were germinated in trays filled with quartz sand in the lab. Ten days after germination, seedlings were transplanted into the microcosms (8 cm in diameter and 12 cm in height) with 250 g of PAHs contaminated soil collected from a chemical plant in Nan**g (118°44′51.87″E, 31°58′4.71″N). Plant communities consisting of nine individuals and differing in functional group richness (8 different communities) were set up: bare ground (no plants); functional group ‘monocultures’ of either three grass species, three herb species, or three legume species; mixtures of two functional groups (grasses plus herbs, grasses plus legumes, or herbs plus legumes); and the mixture containing all three plant functional groups (grasses plus herbs plus legumes), thereby yielding functional group richness levels of 0, 1, 2, and 3 and functionally dissimilar plant communities. Each plant community was replicated four times per CO2 × O3 treatment (32 microcosms per OTC, 128 microcosms in total). Plant communities were cultivated in the lab for one week, and dead seedlings were replaced before microcosms were transferred to OTCs.

The microcosms were randomly placed in the OTCs, and each microcosm was watered with 10–20 ml of distilled water per day. After 10 weeks of cultivation in OTCs, plants and soils were sampled, survival of plants and plant community biomass was measured, and soils for PAHs determination were stored at −20 °C, whereas soils for the measurement of microbial parameters were stored at 4 °C.

Determination of soil enzymatic activity

A very important step of PAH metabolism by bacteria and fungi is the breaking of PAH rings by phenol oxidase or polyphenol oxidase65,66. Therefore, these enzymes were used as proxy for general microbial processes linked to PAH degradation. For enzyme measurements, 0.5 g fresh soil was mixed with 20 ml milli-Q-water in 50 ml falcon tubes, shaken at 250 rpm for 30 min, centrifuged at 3000 rpm for 10 min, supernatants mixed with substrates and buffer in 96-well plates (Corning 96 Flat Bottom Transparent Polystyrol), then determined on a plate reader (Infinite M200, Tecan, Germany). Phenol oxidase activity was measured according to a modified protocol68. Briefly, 20 μl soil supernatant was mixed with 100 μl 5 mM bicarbonate buffer and 100 μl 5 mM L-3,4-dihydroxyphenylalanine (L-DOPA) solution, incubated at 27 °C, and absorbance was measured at 460 nm for 1 h. ΔA460/min from the initial linear portion of the curve was calculated. Polyphenol oxidase activity was measured according to Montgomery and Sgarbieri69. Briefly, 20 μl soil supernatants was mixed with 100 μl 0.5 M potassium phosphate buffer and 100 μl 1 mM 3-(4-hydroxyphenyl) alanine (L-Tyrosine) solution, incubated at 25 °C, and absorbance was measured at 280 nm for 12 min. ΔA280/min from the initial linear portion of the curve was calculated.

PLFA analysis

PLFAs were extracted according to Bligh and Dyer70 modified by Kramer and Gleixner71. Briefly, soil lipids were extracted by a mixture of chloroform, methanol, and 0.05 M phosphate buffer (pH 7.4) and split up into phospholipids by eluting with chloroform, acetone, and methanol from a silica-filled solid phase extraction column. Subsequently, the phospholipids were hydrolyzed and methylated by a methanolic KOH solution, and the PLFA-methyl esters were identified and quantified by GC-ECD (PerkinElmer, Clarus 500, USA). PLFA 19:0 was used as internal standard. Separated phospholipid fatty acid methyl-esters were identified by chromatographic retention time and mass spectral comparison with a mixture of standard qualitative bacterial acid methyl-ester that ranged from C11 to C24 (Supelco). For each sample, the abundance of individual phospholipid fatty acid methyl-esters was expressed in nmol per g dry soil. The nomenclature for PLFAs followed that of Frostegård et al.72. The sum of PLFAs i14:0 i15:0, a15:0, i16:0, i17:0, and i18:0 represented the biomass of Gram-positive bacteria, that of PLFAs cy17:0 and cy19:0 represented the biomass of Gram-negative bacteria, and the amount of the fungal-specific fatty acid 18:2ω6,9 was used as an indicator of fungal biomass73,74.

Determination of PAHs in soils

Soil samples stored at −20 °C were freeze-dried (Labconco 12 L, Labconco Co., USA) for 96 h, ground by mortars, and passed through a 2 mm sieve. Samples (5 g) were extracted with 20 mL methanol: methylene dichloride (1:2, v-v), concentrated in a rotary evaporator, and dried under a fine stream of nitrogen. The residues were dissolved in 0.5 ml acetonitrile. Samples were analyzed by high-performance liquid chromatography on a SpuelcosilTM LC-PAH column (250 × 4.6 mm, 5 μm) (Supelco, Bellefonete, PA, USA) with UV detector at 254 nm (HPLC-UV, Hitachi L2000). The temperature of the column was kept constant at 30 °C to obtain reproducible retention times. The mobile phase consisted of water and acetonitrile in gradient mode at flow rate of 1 ml/min. The gradient solvent system started with 60% acetonitrile in water (v/v) during 10 min, then increasing linearly to 100% acetonitrile within 10 min, the 100% acetonitrile was maintained for 20 min, and finally returned to the initial conditions in 2 min.

Statistical analyses

Analyses of variance (ANOVAs) were performed to test effects of CO2 (ambient and elevated), O3 (ambient and elevated), plant functional group richness (1, 2, 3 functional groups present), and all interactions on total plant biomass, plant shoot biomass, plant root biomass, plant survival, biomass of Gram-positive bacteria, biomass of Gram-negative bacteria, biomass of fungi, phenol oxidase activity, polyphenol oxidase activity, and total PAH residuals (for the latter, treatments with 0, 1, 2, and 3 functional groups were considered). If significant treatment effects were detected, additional Tukey’s HSD tests were performed to test for differences among means. ANOVAs were performed using Statistica 7.1 (Statsoft). Furthermore, structural equation modeling (SEM) was used to shed light on the mechanisms of PAH degradation by accounting for multiple potentially correlated effect pathways to disentangle the direct and indirect effects75 of experimental treatments and soil microbial community properties. The initial model was based on previous knowledge with experimental treatments as exogenous variables and the endogenous variables “Gram-negative bacteria”, “Gram-positive bacteria”, “PC1 microbial PLFAs” (representing PLFA composition), “PC1 PAHs” (representing PAH composition), and “total PAHs”. The adequacy of the models was determined via chi²-tests, AIC, and RMSEA76. Model modification indices and stepwise removal of non-significant relationships were used to improve the models; however, only scientifically sound relationships were considered75. Structural equation modeling was performed using Amos 5 (Amos Development Corporation, Crawfordville, FL, USA).

Data availability

All data generated and/or analyzed during the current study are available from the corresponding author on reasonable request.