Expanding the plant economics spectrum with root nitrogen reallocation

Description

Notes

Methods

A global study for reallocation of N from roots

To position RNR in the root economics space, we compiled a global dataset on N reallocation by searching the keywords of 'nitrogen reallocation' OR 'nitrogen remobilization' OR 'nitrogen redistribution' OR 'root nitrogen' using "Web of Science", "Scopus" and "Google Scholar". Based on the criteria of RNR quantification, only 15 papers met the merit for inclusion. These selected studies applied either ¹⁵N isotope or N budget approaches to quantify RNR (14 versus one study). For the ¹⁵N isotope approach, soil available N pool was labelled with ¹⁵N tracers directly after defoliation, and thereafter the ¹⁵N enrichment of the soil available N pool was used to quantify N uptake from the soil and N remobilization was then estimated as the difference between the N pool in regrown shoots and soil N uptake (Yang et al., 2023). Regarding the N-budget approach, it was also implemented with the facilitation of ¹⁵N tracers, where the percentage of N uptake from soil (i.e., 100%-RNR%) in the second-year growth was equal to the annual depletion rate of ¹⁵N in plant biomass since labelling at the first year (Weinbaum & Kessel, 1998).

¹⁵N isotope labelling

In late July of 2021, a 1 × 1 m subplot was randomly implemented within each treatment plot for tracer ¹⁵N application at peak plant community biomass. Aboveground plant biomass was clipped to 10 mm above the soil surface for each subplot to ensure plant regrowth. Afterwards, the ¹⁵N tracer (99.16 atom%, as ¹⁵NH₄¹⁵NO₃) was dissolved in 500-mL deionized water and uniformly sprayed onto each subplot with a small sprayer. Based on ¹⁵N isotopic signatures, similar to the methodology used in the studies from the above data synthesis, we estimated the contribution of direct N uptake from soil and the contribution of RNR (N stored in roots prior to clipping and reallocated to aboveground organs during their growth). An additional 500 mL of deionized water was sprayed to wash any ¹⁵N attached to plant stubbles down to the soil surface. The tracer application rate was 50 mg ¹⁵N m^-2, which ensured the detection of ¹⁵N signatures without interfering with the natural N cycling processes (Templer et al., 2012). The same tracer application procedure was repeated in late July of 2022, by choosing another subplot of 1 m × 1 m at least 1-m away from the previously labelled subplot within each plot.

Plant and soil sampling

Plant samples clipped before the tracer application were separated into species and dried at 65 °C for 48 h to constant weight. Three weeks after tracer application and plant regrowth, aboveground plant biomass was clipped again to determine the biomass regrowth of each species in the same way (see details in Zhang et al., 2022). We combined and homogenized the species belonging to the same functional group and then ground them into powder for chemical analyses. The ¹⁵N signature was measured on aboveground tissues for each functional group separately.

Unlabelled soils were also collected for background ¹⁵N measurement in the subplots prior to the tracer application. Briefly, topsoil samples (0-10 cm) were collected by randomly selecting three cores and then mixed thoroughly in each subplot. Twenty-one days after labelling, topsoil was sampled again in the same way. Both unlabelled and labelled soil samples were sieved through a 2-mm mesh size to remove the stones and plant residues. Plant and soil samples were collected and processed in the same way for 2021 and 2022.

Given that sorting roots to species or functional group level would be extremely challenging, we chose to quantify root traits only for the most dominant plant species from each functional group. Leymus chinensis (Trin.) Tzvel (L.c), Carex duriuscula C.A.Mey (C.d) and Pulsatilla turczaninovii Kryl. et Serg (P.t) were the dominant species for grasses, sedges and forbs, respectively, accouting for the largest percentages of overall biomass at the community level (68%, 11%, and 6%, respectively; Supplementary Fig. 2a) and in their corresponding functional groups (97%, 100%, and 31%, respectively; Supplementary Fig. 2b-d). Specifically, plant roots were collected by a spade to a depth of 10 cm from a soil block (20 cm long × 15 cm width) around the selected individual plant species and soils adhered to the roots were carefully removed. Root samples were rinsed with distilled water and then frozen at -20 °C for later analysis of morphological traits and AMF colonization.