Three types of biochar were assessed as P sources in P. sylvestris seedling establishment alongside mineral P fertilizer. The response of P. sylvestris root systems to the presence of three biochars of increasing P content and availability was assessed for low-P conditions using rhizoboxes (Fig. 1) and a bespoke growing medium (Table 1b). Root system response to the biochars was compared to their response to mineral P fertilizer.
Biochar Production And Properties
The three biochars were low in P (“Reference biochar”), naturally high in P (“VCZ biochar”), and artificially enhanced in P (“Processed biochar”). All three biochars used were derived from woody biomass feedstock and produced by the UK Biochar Research Centre (UKBRC) (University of Edinburgh, Edinburgh, UK). The equipment used was the Stage III system, a horizontal continuous feed pyrolysis kiln, manufactured by Ansac Pvt Ltd (Bunbury, Australia). The feedstock was pyrolyzed at a nominal highest heating temperature (HTT) of 550 ± 5°C with a heating rate of 78°C min−1. Mean residence time in the kiln was 12 min, with 3.9 min at HTT. Details of the Stage III pyrolysis system can be found in Mašek et al. (2018).
Feedstock for VCZ biochar comprised 40% wood and 60% bark by volume, incorporating the vascular cambium zone of Picea sitchensis (Sitka spruce) and naturally enriched in P (Rathnayake et al. 2021). It is a co-product of debarking during sawmill processing of timber logs and was obtained from the Petersmuir sawmill in south-east Scotland (BSW Ltd, Scotland).
Reference biochar was produced from mixed softwood fuel pellets (Puffin Wood Fuels, Insch, Scotland) using the Stage III pyrolysis described above. The sample represents the standard biochar SWP550 that is available from the UKBRC and widely adopted by the research community (Mašek et al. 2018). For use in this study, the SWP550 biochar was infused in deionized water for 72 hours and fully dried
Processed biochar was created by exposing Reference biochar to dissolved P at a concentration of 410 g total P l−1 with TSP fertilizer (90% water soluble P). The infusion was undertaken at a volume ratio of biochar in P solution and period of 72 hours and fully dried. The P solution was prepared using commercial TSP fertilizer (Progreen Ltd, Bourne, England).
The composition and properties of the three biochars and TSP fertilizer are shown alongside those of the growing medium used in the rhizoboxes in Table 1.
Table 1 Key chemical properties of (a) supply treatment materials and (b) growing medium/treatment mix in rhizobox treatment areas. Values are mean ± SD, n = 3 in a and b except for the Blank, n = 5.
a)
Treatments
|
VCZ biochar
|
Reference biochar
|
Processed biochar
|
TSP fertilizer
|
pH
|
10.0 ± 0.01
|
7.42 ± 0.02
|
3.60 ± 0.02
|
2.74 ± 0.40
|
NH4+-N (mg g−1)
|
< 0.01
|
< 0.01
|
0.401 ± 0.09
|
1.12 ± 0.11
|
NO3−-N (mg g−1)
|
< 0.02
|
< 0.02
|
0.08 ± 0.01
|
0.722 ± 0.050
|
Available P (mg g−1)
|
2.24 ± 0.84
|
0.542 ± 0.010
|
258 ± 6
|
602 ± 3
|
Fe (mg kg−1)
|
22.9 ± 11.3
|
66.4 ± 5.7
|
29.1 ± 0.5
|
189 ± 3
|
Al (mg kg−1)
|
21.4 ± 10.4
|
42.4 ± 3.6
|
148 ± 8
|
387 ± 6
|
Ca (g kg−1)
|
12.4 ± 5.0
|
2.71 ± 0.05
|
19.7 ± 0.5
|
53.2 ± 0.9
|
Cd (mg kg−1)
|
0.06 ± 0.01
|
0.05 ± 0.01
|
1.49 ± 0.04
|
7.61 ± 0.20
|
b)
Treatments
|
Blank
|
VCZ biochar
|
Reference biochar
|
Processed biochar
|
TSP fertilizer
|
pH
|
4.41 ± 0.02c
|
8.12 ± 0.01a
|
5.58 ± 0.01b
|
3.65 ± 0.02d
|
2.9 ± 0.02e
|
NH4+-N (mg g−1)
|
1.41 ± 0.06a
|
0.798 ± 0.047b
|
0.742 ± 0.031b
|
0.754 ± 0.087b
|
1.12 ± 0.11a
|
NO3−-N (mg g−1)
|
1.04 ± 0.02a
|
0.654 ± 0.029b
|
0.423 ± 0.044c
|
0.201 ± 0.009d
|
0.722 ± 0.053b
|
Available P (mg g−1)
|
0.046 ± 0.020e
|
1.24 ± 0.86c
|
0.314 ± 0.048d
|
156 ± 24b
|
590 ± 37a
|
Fe (mg kg−1)
|
64.9 ± 3.0b
|
20.8 ± 1.2c
|
8.91 ± 0.23d
|
23.5 ± 1.2c
|
173 ± 4a
|
Al (mg kg−1)
|
9.52 ± 0.54d
|
15.6 ± 0.5c
|
11.2 ± 0.4d
|
107 ± 8b
|
365 ± 2a
|
Ca (g kg−1)
|
0.135 ± 0.007e
|
4.72 ± 0.19c
|
1.21 ± 0.03d
|
11.2 ± 0.7b
|
50.2 ± 1.0a
|
Cd (mg kg−1)
|
0.04 ± 0.01c
|
0.06 ± 0.01c
|
0.05 ± 0.01c
|
1.11 ± 0.04b
|
7.12 ± 0.33a
|
* Different letters indicate significant differences at p < 0.05 identified from One-way ANOVA. |
Rhizobox Experiment Design
Rhizoboxes were made of 0.6 cm thickness PVC sheet. The effective size was 40 cm height x 30 cm width x 0.6 cm thick (Fig. 1). The separation was customized for one-year-old P. sylvestris seedlings, considering the diameter of the stem and the length and density of their root systems. The rhizobox comprised five parts in total including the sides and base, which had machined groves along joins to provide additional strength and prevent potential slippage. The transparent front sheet was detachable and designed for observation and photographic imaging. Black opaque sheets were used for the other sides and were fixed along the exterior of the joins using polyurethane grab adhesive (Gorilla Glue Co., Sharonville, Ohio). Rhizoboxes were set for 24 hours and designed to prevent ingress of light and maintain consistent conditions inside and outside the rhizoboxes. Following the completion of assembling, they were rinsed, alcohol-sterilized and dried before use.
Bespoke growing medium was prepared and sterilized 2 weeks prior to the set-up. It comprised of 60% peat and 40% quartz sand by volume, which ensured key nutrients were initially at deficient level (Table 1a). The growing medium had no nutrients added and displayed nutrient status comparable to particularly poor forest soil (Pyatt et al. 2001), accurately reflecting P-limited plantation forests in the UK. The growing medium was passed through a 2mm sieve prior to filling the prepared rhizoboxes. Approximately 720 g of growing medium was added into each rhizobox. The variation of added growing medium was no more than 10% to maintain a bulk density of 1 ± 0.1 kg m−3 to ensure a comparable growing environment from the outset. If the measured pH (McLean 1983) of growing medium was outside the range 4.5 ± 0.5 it was adjusted using 0.01 mol l−1 NaOH solution and 0.01 mol l−1 H2SO4. The growing medium in the rhizoboxes was fully watered with deionized water and stabilized for 48 hours before the seedlings were planted.
The 30 cm x 40 cm frontal area of the rhizoboxes were divided into twelve unit-sectors with a size of 10 cm width x 10 cm height which were marked onto the front sheet to create a visual grid. The four treatment materials were applied at differential mass dose after the growing medium was fully stabilized based on the recommended dose of P fertilization for conifer seedlings in P-deficient soils (Taylor 1991) (adjusted for targets application and growing medium volume, to maintain equivalent TSP : growing medium by volume): 0.96 g TSP fertilizer, 4.56 g Processed biochar. 3.60 g Reference biochar and 3.60 g VCZ biochar. All biochar treatments had equal mass dose of biochar, with processed biochar having additional biochar due to TSP infusion of equal dry weight of TSP to TSP treatment. The treatments were applied centrally in the lower left sector without any mesh exclusion. The treatment area (TA) for each treatment was 2 cm, 4 cm, 5.5 cm and 7.8 cm, respectively using prepared paper templates. The same volume of growing medium (1:1) was introduced to ensure the bulk density approximately the same between different treatments. Each treatment was arranged with five replicates. A blank control (“Blank”) comprising growing medium was included, resulting in 25 rhizoboxes in total.
One-year-old container grown P. sylvestris seedlings were obtained from Forest Research (Northern Research Station, Roslin, Scotland) as P. sylvestris is the most tolerant and suitable conifer species in sandy P-deficient soils with highest commercial profits. These trees had been grown from seed and all seedlings were in a dormant state. The seedlings had all been grown using the same growing medium and all displayed root systems 20 - 30 cm in length and were not nutrient deficient. Seedlings were carefully removed from containers and growing medium was manually removed from the root systems. They were carefully rinsed to avoid transfer of nutrients from the original growing medium. The seedlings were planted into the rhizoboxes approximately at a central position, gently separating entangled roots. To diminish the possibility of ‘transplant shock’, roots were not sterilized or trimmed. To ensure roots could consistently detect those P sources, one root in each rhizobox was positioned 2 cm adjacent to the edge of the TA, oriented towards the center. Each rhizobox was watered weekly by removing the front sheet and evenly spraying the surface medium with distilled water using a mist sprayer. The watering regime was flexible to ensure that the growing medium was moist but with no excess water or vertical water movement or pooling.
The front (colorless) PVC sheets were clipped to the treated, planted boxes. Polystyrene sheets cut to the same size and shape were used to exclude light and maintain consistent internal conditions. They were also used to loosely cover the surface of the growing medium to reduce soil surface light penetration and temperature gradients, while ensuring respiration was not impeded. The temperature and relative humidity in the glasshouse was between 17.0 - 27.2°C and 36.5 - 100.0%, respectively during the 10 weeks experimental period. Due to space limited, 10 weeks later, interior surfaces were fully covered by roots and new roots started to generate towards deeper growing medium.
The experiment used a randomized block design, with one replicate of each treatment randomly placed in each block. The rhizoboxes in each block were stacked together at an angle of 60°, taking care to ensure the branches of above-ground parts did not interfere and were horizontal and vertically staggered (Fig. 1).
Morphological Monitoring And Analysis Of Root Growth
The root system in each rhizobox was imaged photographically once each week. A lightbox was constructed from PVC sheet enveloped with thick photographic curtain. A port was created at the top of the box to provide access for a camera lens. The box was illuminated internally with LED symmetrical lighting strips orientated in four evenly distributed parallel lines across the top of the box, while thick black felt was used to line the base and sides to eliminate the ingress of natural light. The rhizoboxes were imaged in turn, with a fixed camera position in the light sealed port and with a precisely determined position for the rhizobox that ensured consistency in image positions. The rhizoboxes were imaged weekly using a SLR camera at a native resolution of 7360 x 4912 pixels.
A method to spatially track and analyze root growth was developed using ArcGIS Pro 2.6 software (ESRI, Redlands, USA). The first step was to map each image to a georeferenced 30 cm x 40 cm physical area (the effective rhizobox area). This provided the geographic coordination required for ArcGIS Pro 2.6 to define absolute position. A feature layer was then created to manually track growth of individual roots, assisted by the association between root color and increasing root age. New roots develop on the interior surface of the PVC owing to geotropism and the availability of water and air. Only new roots growing on the interior surface were taken into account.
Weekly images were analyzed as separate segments for the same rhizobox image. Overlaying the images in this way allowed an integrated root growth feature layer to be created and summarized for each rhizobox. This provided the calculated length and density of cumulative root growth. The analysis also allowed length, density and growth rate to be visualized and analyzed. Root growth was separately assessed for the whole rhizobox, TA and TA3.
Sampling And Chemical Analysis
10 weeks later, the root system was separated from the seedling at the first root branch and was gently rinsed, dried in an oven at 90°C and weighed on an electronic balance (accurate to 0.001 g). New needles were picked individually from the top and oven dried at 90°C before further total P analysis.
Growing medium, treatment materials and treatment area (mix of treatment materials and growing medium, TA), were sampled at the beginning before the seedlings were introduced and were pooled across replicates for chemical analysis (Table 1b). After 10 weeks the TA and the growing medium in each rhizobox were separately sampled, using prepared paper templates to ensure precise positioning. The material sampled within each area was then thoroughly mixed to reduce intra-sample area variation and transferred to a refrigerator below 4°C. Samples were oven dried at 90°C and were ground in a ball grinder before further analysis.
Chemical analysis was undertaken to determine available P and total P, Fe, Al, Ca and other heavy metals to elucidate potential factors influencing root-biochar interactions. Extraction or digestion procedure for each sample was replicated for 3 times. Available P of samples was analyzed through water extraction (Prendergast-Miller et al. 2014) and subsequent analysis of the extracts by autoanalyzer (Bran Luebbe AA3, Seal Analytical, Norderstedt, Germany). Total P, Fe, Al, Ca and other heavy metals were extracted by a modified dry-ash method for biochar and measured by inductively coupled plasma optical emission spectrometry (ICP-OES, Thermo-iCAP 6300, Thermo Electron, Waltham, USA). Samples were heated in a muffle furnace (500°C, 8 h) and placed into a steam bath upon cooling. Concentrated nitric acid (HNO3) was added and evaporated to dryness, followed by 1:4 HNO3 and H2O2 which was evaporated to dryness. All residues were transferred through Whatman Grade 4 filter paper, using HNO3 and deionized water, into a volumetric flask. The elemental composition of the filtrate was analyzed using inductively ICP-OES. The pH of growing medium was assessed for the zone within 3 cm of treatment area (TA3). This was done after 10 weeks plant growth using a soil : water ratio of 1:2 (McLean 1983).
Data analysis
ArcGIS Pro 2.6 software was used for initial length and density calculation of new roots and then transferred to point features for kernel density analysis which can reflect the density of roots in the neighborhood of each output raster cell. pH data was tested for normality (K-S test) and homogeneity of variance (Levene test, p = 0.05). One-way analysis of variance (ANOVA) was conducted using Python 3.8 and with p = 0.05 as the threshold for statistical significance. Linear and nonlinear LangmuirEXT1 regression models were developed using Origin Pro 2018 statistical software (Origin Lab, Northampton, USA).