Switchgrass (Panicum virgatum L.), as a dedicated bioenergy crop, can provide cellulosic feedstock for biofuel production while improving or maintaining soil quality. However, comprehensive evaluations of how switchgrass cultivation and nitrogen (N) management impact soil and plant parameters remain incomplete. We conducted field trials in three years (2016–2018) at six locations in the North Central Great Lakes Region to evaluate the effects of cropping systems (switchgrass, restored prairie, undisturbed control) and N rates (0, 56 kg N ha^-1 yr^-1) on biomass yield and soil physicochemical, microbial, and enzymatic parameters. Switchgrass cropping system yielded an aboveground biomass 2.9–3.3 times higher than the other two systems (Jayawardena et al., In submission) but our study found that this biomass accumulation didn’t reduce soil dissolved organic C (DOC), total dissolved N (TDN), or bacterial diversity. The annual aboveground biomass removal for bioenergy feedstock, however, reduced soil microbial biomass C (MBC) and N (MBN) and bacterial richness in the 2^nd and 3^rd years; despite this, continuous monocropping of switchgrass improved soil TDN, inorganic N, bacterial diversity, and shoot biomass in the 2^nd and/or 3^rd years when compared to the 1^st year. N fertilization increased aboveground biomass yield by 1.2 times and significantly increased soil TDN, MBN, and the shoot biomass of switchgrass when compared to the unfertilized control. Locations with higher C and N contents and lower C:N ratio had higher aboveground biomass, MBC, MBN, and the activity of BG, CBH, and UREA enzymes; by contrast, locations with higher pH had higher soil TDN and activity of NAG and LAP enzymes. Our research demonstrates that switchgrass cultivation could improve or maintain soil N content and N fertilization can increase plant biomass yield. The comprehensive data also can inform future biogeochemical models to successfully implement switchgrass for bioenergy production.

Soil physicochemical properties determination

Dissolved organic carbon (DOC) is a component of the soil active organic carbon (Matlou and Haynes, 2006), which is an organic carbon source for microorganisms in the soil. In this study, soil carbon (C) and nitrogen (N) in the labile organic pools (dissolved organic C, total dissolved N), inorganic pools (NH4⁺, NO3^-, plant-available N), microbial pools (microbial biomass C, microbial biomass N), as well as soil pH and soil moisture were determined.

Six grams of soil were extracted by 30 ml of 0.5 M K2SO4 and filtered with Whatman® filter paper (Grade 202) as described in Smercina et al. (2021). Dissolved organic C (DOC) and total dissolved N (TDN) were determined using a vario TOC cube (Elementar Americas, Inc., Ronkonkoma, NY, USA) following the manufacturer’s instruction. Soil inorganic N (NH4⁺, NO3^-) in the extractant was determined using 96-well high-throughput colorimetric methods as described by Smercina et al. (2021). Soil microbial biomass C (MBC) and N (MBN) were determined by the chloroform fumigation direct extraction method as described by Anderson and Domsch (1978) and Gregorich et al. (1990) and a vario TOC cube (Elementar Americas, Inc., Ronkonkoma, NY, USA)  following the manufacturer’s manual. MBC and MBN were calculated using DOC and TDN contents according to the equations reported by Beck et al. (1997) and Brookes et al. (1985). Soil pH was determined in a 1:2 (w/v) soil: deionized water extractant with a VWR Symphony B20PI Benchtop pH meter (VWR International, LLC., Radnor, PA, USA) (Schofield and Taylor, 1955). Soil moisture was measured following the gravimetric method described by Reynolds (1970).

Soil microbial richness and diversity assessment

Soil microorganisms play a critical role in organic matter decomposition, nutrient cycling, and soil productivity (Ramírez et al., 2020; Fierer et al., 2021). Soil DNA for 2016 and 2017 was extracted from 0.5 g soil using a PowerSoil® DNA Isolation Kit (MO BIO Laboratories, Inc., Carlsbad, CA, USA) following the manufacturer’s protocol. Soil DNA for 2018 was extracted in 96-well plates using the KingFisher Flex Purification System (Thermo Fisher Scientific, Waltham, MA, USA) with the PowerSoil® kit. The extracted soil DNA was electrophoresed on 1% agarose gels, and the quality and quantity of DNA were evaluated using a NanoDrop-1000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). For bacteria, the V4 hypervariable region of 16S rRNA genes was amplified using 515F/806R primers (Caporaso et al., 2011), and the Illumina compatible libraries were prepared using primers containing both the target sequences and the dual indexed Illumina compatible adapters (Kozich et al., 2013) by Michigan State University (MSU) Research Technology Support Facility (RTSF) Genomics core. For fungi, ITS1 region was amplified using ITS1-F/ITS2 primers (White et al., 1990), and libraries were multiplexed using a three-step PCR sequence as described by Chen et al. (2018). The completed libraries were normalized using Invitrogen SequalPrep DNA Normalization plates and pooled and cleaned up using AmpureXP magnetic beads. Libraries were then paired-end sequenced by MSU RTSF Genomics core on a MiSeq platform (Illumina Inc., USA) using the v2 kit for bacterial libraries and the v3 kit for fungal libraries. Bioinformatics and sequence processing was conducted using Quantitative Insights Into Microbial Ecology (QIIME) 2 (Bolyen et al., 2019) and USEARCH (Edgar, 2010), and details can be found in the supplementary material.  A total of 21,450 bacterial OTUs and 2,824 fungal OTUs were rarefied to 10,000 reads to evaluate the two richness indices, Chao1 (Chao, 1984) and Abundance-based Coverage Estimator (ACE) (Chao and Lee, 1992), and two diversity indices, Shannon-Weiner (Shannon, 1948) and reverse Simpson (Invsimpson) (Simpson,1949). All diversity metrics were calculated using the vegan package in R (Oksanen et al., 2014).

Soil carbon (C) and nitrogen (N) cycling-related enzymes activity assay

Soil enzyme activities are the indicators of microbial community and functions. Soil extracellular enzymes decompose substrates of varying composition and complexity (Sinsabaugh, 2010; Jian et al., 2016) and play an important role in C sequestration and N cycling (Bowles et al., 2014; Keane et al., 2020). In this study, five C and N cycling-related enzymes, cellobiohydrolase (CBH, also called β-d-cellobiosidase) and β-glucosidase (BG), urease (UREA), N-acetyl-β-glucosaminidase (NAG), and leucine aminopeptidase (LAP), were determined to evaluate how switchgrass cultivation and N rate affect soil C sequestration and N availability.

Soil β-glucosidase (BG, EC 3.2.1.21) and cellobiohydrolase (CBH, EC 3.2.1.91) are commonly studied extracellular glycosidases to reveal the potential microbial activities associated with fast-turnover organic C (Klose and Tabatabai, 2002; Sinsabaugh et al., 2008). Soil BG catalyzes the hydrolysis of β-D-glucopyranoside and is involved in the saccharification of cellulose (Deng and Tabatabai, 1994; Tabatabai et al., 1994; Bandick and Dick, 1999; Turner et al., 2002); CBH hydrolyzes the end of the cellulose chain and produces glucose or cellobiose as the end product (Lynd et al., 2002). Of the three N-cycling enzymes, urease (UREA, EC 3.5.1.5) regulates the soil N transformation and is in charge of the hydrolysis of urea into ammonia and CO₂ (Kong et al., 2008); soil N-acetyl-β-glucosaminidase (NAG, EC 3.2. 1.30) and leucine aminopeptidase (LAP, EC 3.4.11.1) regulate the release of plant-available N from organic compounds (Sinsabaugh et al., 2008). Activities of NAG, BG, CBH, and LAP were measured by a fluorometric method (Saiya-Cork et al., 2002; DeForest, 2009; German et al., 2012; Kim et al., 2018) and a BioTek microplate reader (BioTek Instruments, Winooski, VT, USA). Activities of NAG, BG, CBH were measured at an excitation wavelength of 370 nm and an emission wavelength of 455 nm, and activity of LAP was determined at an excitation wavelength of 350 nm and an emission wavelength of 430 nm. Their activities were expressed as nmol g⁻¹ dry soil h⁻¹. Soil urease (UREA) activity was measured using the method described by Sinsabaugh et al. (2000) and urea (Millipore Sigma, 57-13-6) and determined spectrophotometrically at 610 nm. The activity of UREA was expressed as nmol NH4⁺ g⁻¹ soil h⁻1.

Determination of plant traits and biomass yield

In July each year, the aboveground plant height, shoot biomass, root biomass, root length, and root width of switchgrass in each plot were determined. Switchgrass roots from the core were washed and scanned with an Epson Perfection V600 Photo Scanner (Seiko Epson Corporation, Suwa, Nagano, Japan), and the 1200-dpi image was compressed to 400-dpi and analyzed with Gia Roots to measure the length and width of the longest roots (Galkovskyi et al., 2012). The shoot and root were oven-dried at 60 oC for at least 48 hours until a constant weight for determining shoot dry biomass and root dry biomass. At field harvest during mid-October and mid-November each year, all plants were weighed on the same day as the fresh biomass. Plant subsamples were collected and dried at 60 °C for at least 48 hours until constant weight for plant moisture determination and dry biomass calculation.