Introduction

Urban greening provides a range of social, health, economic and environmental benefits that improve the liveability of cities and human well-being (Bolund and Hunhammar 1999; Tzoulas et al. 2007; Salmond et al. 2016; Palliwoda et al. 2020; Marselle et al. 2020). For this reason, investment in urban greening is growing globally (Pincetl et al. 2013; Mell 2016). For example, several Australian state- and local-level governments have adopted ambitious canopy cover targets for their urban areas (Department of Environment, Land, Water and Planning 2017; Greater Sydney Commission 2018). However, urban areas are often challenging environments for plants to survive and thrive in because of space limitations, compacted soils, water and nutrient limitation and reduced soil microbial activity (Gilbertson and Bradshaw 1985; Clark and Kjelgren 1990; Haaland and van den Bosch 2015; Ferreira et al. 2018). Therefore, nursery and urban forest practitioners need to implement practices that enhance plant establishment in urban areas to ensure strong urban greening outcomes in the future.

One practice that is widely used by nursery and urban forest practitioners to enhance plant establishment is root pruning (e.g. butterflying, shaving, vertical sliding) before transplanting (Geisler and Ferree; Allen et al. 2017). When pruned, roots form a meristematic callus at the injury site, which reduces root defects (e.g. kinking, spiralling) and stimulates lateral root growth. This in turn increases the volume of soil reached by the roots, which improves anchorage as well as water and nutrient uptake (Geisler and Ferree 2011). However, harsher root pruning methods (e.g. trenching, pruning after bare rooting) may cause stress to the plant because the roots are not able to maintain the water and nutrient uptake necessary to support their living biomass (Wilson 1988; Hamilton 1989). As a result, these methods may reduce shoot growth in plants in the short-term (Watson 1998; Andersen et al. 2000; Fini et al. 2013; Benson et al. 2019), particularly when plants are transplanted from the nursery to the field, where conditions tend to be harsher (Close et al. 2005). Milder root pruning methods (e.g. root shaving), on the other hand, are less likely to have detrimental impacts on shoot growth (Gilman et al. 2010; Gilman 2013; Cregg and Ellison 2018). Nevertheless, it is important that we implement practices to help mitigate root pruning stress when it occurs to ensure the best nursery production and urban greening outcomes possible.

Biostimulants are a diverse range of extracts obtained from organic raw materials that contain bioactive compounds (Bulgari et al. 2015). Biostimulants can have a range of benefits for plant growth, nutrition and stress resilience (Van Oosten et al. 2017; Baltazar et al. 2021; Pereira et al. 2021; Li et al. 2022). Biostimulants may mitigate root pruning stress by enhancing anti-oxidant activity, nutrient uptake efficiency, stress resistance gene expression, stress signalling compound production and water use efficiency (García-García et al. 2020; Ma et al. 2022). However, the mode of action of many biostimulants is complex and not well understood (Baltazar et al. 2021). Some of the most common biostimulants are derived from humic substances, protein hydrolysate and seaweed extract (Calvo et al. 2014; Du Jardin 2015). Humic substance biostimulants interact with plant membrane transporters responsible for nutrient uptake and membrane-associated signal transduction cascades that regulate growth and development (Canellas et al. 2015). Protein hydrolysate biostimulants stimulate C and N metabolism, as well as regulate the activity of enzymes involved in N assimilation and the tricarboxylic acid cycle (Colla et al. 2017). Seaweed extract biostimulants initiate the production of reactive oxygen and nitrogen species, and NADPH oxidase, which enables enhanced growth regulation by the plant (Ali et al. 2021) Research on the use of biostimulants in mitigating root pruning stress has primarily focused on edible and ornamental plant species (Colla and Rouphael 2015; Li et al. 2022). Although a recent survey of urban forestry practitioners in Australia revealed the main reason they use biostimulants is to improve transplant success (Cinantya et al. 2023), little is known about the effectiveness of biostimulants in reducing root pruning stress in urban plant species. Further, there is no clear pattern in the effect of biostimulants on urban plant species performance in general, with both positive (Ferrini and Nicese 2002; Fraser and Percival 2003) and no effects (Fraser and Percival 2003; Abbey and Rathier 2005; Percival 2006; Banks and Percival 2012, 2014) being reported.

This study aimed to determine 1) the short- and long-term impacts of root shaving, a type of root pruning, on the growth of urban plant species, and 2) whether biostimulant application mitigates these impacts. To address these aims, we applied root shaving (not shaved, shaved) and biostimulant (control, humic substance, protein hydrolysate, seaweed extract) treatments to six tree species that are commonly planted in the Sydney metropolitan area, Australia in a factorial design. The study consisted of a glasshouse experiment to simulate nursery production conditions and a field experiment to simulate urban field conditions. The establishment of the study species after treatment was assessed by measuring their assimilation rate, growth and biomass allocation. We hypothesised that:

  1. 1.

    Root shaving would reduce shoot growth in the short-term (16 weeks) but not in the long-term (> 1 year).

  2. 2.

    Biostimulant application would mitigate the impact root shaving has on shoot growth.

Materials and method

Species selection

For the glasshouse experiment, we selected six tree species that are commonly planted in the Sydney metropolitan area, Australia and are typically root pruned, often by shaving, before transplanting (Table 1). For the field experiment, we selected three of the tree species that were used in the glasshouse experiment (Table 1). We selected the study species based on their propagation popularity (i.e. number of Australian growers that propagate them; Table 1; Plummer unpublished) and stock availability from a commercial nursery (Andreasens Green, Kemps Creek, NSW, Australia). Seedlings of each species were propagated and grown in their containers in the same shade house by the commercial nursery for ~20 weeks before the experiment commenced.

Table 1 The taxonomy, number of growers in Australia and container size of the six tree species selected for the study
Full size table

Experimental design

Glasshouse study

The glasshouse experiment was conducted in the Macquarie University Plant Growth Facility (33.775°S, 151.117°E). It was fully factorial with two treatments: root shaving (not shaved, shaved) and biostimulant (control, seaweed extract, humic substance, protein hydrolysate). This design resulted in eight treatment groups. Each treatment group had ten replicates per species, resulting in a total of 480 plants (6 species × 8 treatments × 10 replicates). In addition, 7 replicates of each species were prepared for pre-treatment harvest (6 species × 7 replicates = 42 plants).

Pre-treatment plant height and stem diameter measurements were taken for each plant. Plant height was measured from the soil surface to the apical meristem, while stem diameter was measured at 15 cm above the soil surface using a calliper. Once the pre-treatment measurements were taken, we destructively harvested a sub-set of plants (i.e. pre-treatment harvest), which were oven-dried at 70 °C for 48 h and weighed using an analytical scale (Mettler Toledo, Port Melbourne, VIC, Australia). For each species, we developed a simple linear regression equation for the dry-weight biomass as a function of plant height. Based on these equations, the biomass of the treatment plants was determined.

The treatment plants were evenly distributed based on plant height into the eight treatment groups to ensure an even size distribution across treatments. For the plants allocated to the root shaving treatments, approximately 1 cm of root material was shaved from around their root ball using a sharp knife (Fig. S1). In contrast, the sides of the root ball of the control plants (i.e. not shaved) were not disturbed. All plants had the base of the root ball lightly teased apart. Immediately after preparation, the plants allocated to receive a biostimulant treatment were immersed in a solution of their designated biostimulant for 5 min, while the control plants were immersed in water. The biostimulants used were: 1) humic substance (5 mL/L; Liquid Humus, Nutri-Tech Solution, Yandina, QLD, Australia; Active ingredient: Potassium humate); 2) protein hydrolysate (5 mL/L; Hydrofish, No Frills Fertilisers, Margaret River, WA, Australia; Active ingredient: Fish waste; 3.1N:0.34P:0.34 K); and 3) seaweed extract (SW; 6.25 mL/L; Seasol, Seasol International, Boronia, VIC, Australia; Active ingredient: Kelp; 0.1N:0.01P:1.5 K).

After immersion, the plants were transplanted into 8 L pots (240 mm height × 250 mm diameter) filled with organic potting mix (Australian Growing Solutions, Tyabb, VIC, Australia), which had a slow-release fertiliser (16N:4.4P:8.3K) homogenised into it. Treatments and species were evenly split among 10 blocks, each containing 48 plants (i.e. 6 species × 8 treatments). These blocks were divided across three glasshouses. The temperature of the glasshouses was set to 25 °C day/21 °C night and was continuously maintained by a fan coil unit using a water cooling and heating system. In addition, relative humidity (glasshouse 1 = 80 ± 0.2%, glasshouse 2 = 73 ± 0.2%, glasshouse 3 = 79 ± 0.2%) and photosynthetically active radiation (glasshouse 1 = 783 ± 35 µmol m−2s−1, glasshouse 2 = 478 ± 21 µmol m−2s−1, glasshouse 3 = 601 ± 26 µmol m−2s−1) were continuously monitored using a MultiGrow controller system (Autogrow Systems, Auckland, New Zealand). To ensure that there was no glasshouse effect, the plants rotated through the glasshouses every 5 weeks, spending ~5 weeks in each glasshouse. The plants were mist-watered using overhead irrigation sprinklers three times a day for 5 min (~2.1 L/plant/day) but received no additional fertiliser. Throughout the experiment, pest and fungal outbreaks were treated as required using tau-fluvalinate (Mavrik; Yates, Padstow, NSW, Australia) and polysulfide sulphur (Lime Sulfur; Yates, Padstow, NSW, Australia) respectively.

Field experiment

The field experiment was conducted in Macquarie University Fauna Park (33.769°S, 151.115°E). We tested the same treatments as the glasshouse experiment but only for three of the study species (E. reticulatus, F. macrocarpa ‘Flash’, S. glomulifera) and seven replicates, resulting in a total of 168 plants (3 species × 8 treatments × 7 replicates).

The root shaving and biostimulant treatments were applied to the plants using the same methods as the glasshouse experiment. After treatment, the plants were planted into wok-shaped holes (20 cm deep × 25 cm wide) that were backfilled with unamended field soil and topped with a ~2 cm layer of sand soil (4:1) mix (Australian Native Landscapes, Terrey Hills, NSW, Australia). A ~5 cm layer of Eucalypt wood chip mulch was added to the soil surface around the base of the plants across the planting site, with 10 cm around the base of each plant left as bare soil. The plants were then supported using bamboo stakes. Treatments and species were evenly split across seven blocks, each containing 24 plants (i.e. 8 treatments × 3 species). Within each block, plants were spaced in a 1 m grid (8 m × 3 m). The plants received no supplementary watering (i.e. precipitation only) or fertiliser. Throughout the experiment, glyphosate (i.e. Roundup; Monsanto, St Louis, MO, USA) was used for weed suppression.

Growth and biomass allocation measurements

Glasshouse experiment

The assimilation rate of eight plants from each treatment × species combination was measured 1, 2, 3, 4, 5 and 10 weeks after treatment. For each plant, the assimilation rate was measured on three randomly selected newly fully expanded outer-canopy leaves using a LICOR-6800 portable photosynthesis machine (LICOR Biosciences, Lincoln, NE, USA). The temperature (25 °C), relative humidity (75%), photosynthetically active radiation (500 µmol m−2s−1) and CO2 concentration (450 ppm) within the machine’s cuvette were kept constant between measurements. The measurements were conducted between 08:30 and 13:30 when the plants were the most photosynthetically active and were blocked over time with eight blocks. Note that in the Results, we only present the five weeks after treatment assimilation rate data because there were no treatment effects over time.

The plant height and stem diameter of each plant were re-measured 16 weeks after treatment. The treatment plants were harvested, separated into roots and shoots, oven-dried at 70 °C for 48 h and weighed using an analytical scale (Mettler Toledo, Port Melbourne, VIC, Australia). To calculate the biomass, plant height and stem diameter growth, the following formula was used:

$$\begin{aligned}Growth&=Post\text{-}treatment\;measurement\\&-Pre\text{-}treatment\;measurement\end{aligned}$$

The root and shoot biomass of each plant from the post-treatment harvest were used to calculate biomass allocation (i.e. root-to-shoot ratio or R:S).

Field experiment

Plant height and stem diameter of each plant were re-measured 16, 52 and 120 weeks after treatment. Plant height and stem diameter growth were calculated using the same formula as the glasshouse experiment.

Statistical analyses

Glasshouse experiment

Two-way ANOVAs were used to determine the effect of the root shaving and biostimulant treatment on the growth and biomass allocation of each study species. The root shaving and biostimulant treatments were fixed factors. The response variables were assimilation rate, plant height growth, stem diameter growth, biomass growth and R:S. Tukey post-hoc comparisons were conducted when significant differences between treatments were found. When the normality and/or equal variance assumptions of ANOVA were violated, even after being log transformed, Kruskal–Wallis non-parametric analyses were conducted. Mann–Whitney post-hoc comparisons were conducted when significant differences between treatments were found.

Field experiment

Two-way ANOVAs were used to determine the effect of the root shaving and biostimulant treatment on the growth of each study species. The root shaving and biostimulant treatments were fixed factors. The response variables were plant height growth and stem diameter growth. Tukey post-hoc comparisons were conducted when significant differences between treatments were found.

All statistical analyses were conducted using R (v. 4.2.2, R Development Core Team 2022), with the significance level set at 0.05.

Results

Glasshouse experiment

Assimilation rate

There was no interaction between the root shaving and biostimulant treatments for the assimilation rate of the study species. However, root shaving reduced the assimilation rate of F. australis (F1,53 = 4.23, P = 0.045) and S. glomulifera (F1,55 = 17.39, P < 0.001) (Fig. 1). The biostimulant treatments did not affect the assimilation rate of the study species.

Fig. 1
figure 1

The median assimilation rate of the glasshouse study species five weeks after treatment for each root shaving treatment. Shaving treatment abbreviations: NS not shaved, S shaved. Each box represents the 50% interquartile range, while the dots represent outliers. Asterisks indicate significant differences (P < 0.05) between the shaving treatments within the study species

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Plant growth

There was no interaction between the root shaving and biostimulant treatments for any of the growth metrics measured for the study species. Similarly, the biostimulant treatments did not affect the growth metrics. However, root shaving reduced the biomass (F1,72 = 6.20, P = 0.015; Fig. 2a) and plant height (F1,72 = 12.99, P = 0.001; Fig. 2b) growth of E. reticulatus, the biomass growth of F. australis (F1,72 = 9.10, P = 0.004; Fig. 2a), the biomass growth (F1,71 = 9.40, P = 0.003; Fig. 2a) and plant height (F1,71 = 5.54, P = 0.021; Fig. 2b) growth of L. confertus and the plant height (F1,71 = 5.54, P = 0.021; Fig. 2b) and stem diameter (H7 = 14.64, P = 0.041; Fig. 2c) growth of S. glomulifera.

Fig. 2
figure 2

The median a biomass, b plant height and c stem diameter growth of the glasshouse study species for each root shaving treatment. Shaving treatment abbreviations: NS not shaved, S shaved. Each box represents the 50% interquartile range, while the dots represent outliers. Asterisks indicate significant differences (P < 0.05) between the shaving treatments within the study species

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Root-to-shoot ratio

There was no interaction between the root shaving and biostimulant treatments for the R:S of the study species. However, root shaving reduced the R:S of H. pendula (F1,72 = 11.06, P = 0.001) but increased the R:S of S. glomulifera (F1,72 = 5.52, P = 0.022) (Fig. 3). The seaweed biostimulant increased the R:S of S. glomulifera compared to the humic substance biostimulant (F3,72 = 3.49, P = 0.020).

Fig. 3
figure 3

The median R:S of the glasshouse study species for each root shaving treatment. Shaving treatment abbreviations: NS not shaved, S shaved. Each box represents the 50% interquartile range, while the dots represent outliers. Asterisks indicate significant differences (P < 0.05) between the shaving treatments within the study species

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Field experiment

Plant growth

There was no interaction between the root shaving and biostimulant treatments for the plant height and stem diameter growth of the study species. However, root shaving reduced the stem diameter growth of E. reticulatus (F1,48 = 4.29, p = 0.044; Fig. 4d) and the plant height (F1,47 = 4.52, p = 0.039; Fig. 4a) and stem diameter (F1,46 = 8.00, p = 0.007; Fig. 4d) growth of S. glomulifera 16 weeks after treatment but not 52 and 120 weeks after treatment. The root shaving treatments did not affect the plant height growth of E. reticulatus and F. macrocarpa. The humic substance biostimulant increased the stem diameter growth of E. reticulatus compared to the protein hydrolysate biostimulant 52 weeks after treatment (F3,48 = 3.49, P = 0.020) but not 16 and 120 weeks after treatment. The biostimulant treatments did not affect the plant height growth of the study species.

Fig. 4
figure 4

The median plant height and stem diameter growth of the field study species a, d 16, b, e 52 and c, f 120 weeks after treatment. Shaving treatment abbreviations: NS not shaved, S shaved. Each box represents the 50% interquartile range, while the dots represent outliers. Asterisks indicate significant differences (P < 0.05) between the shaving treatments within the study species

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Discussion

Root shaving before transplanting improves root structure, which enables the plant to explore the soil for resources while providing better support (Gilman et al. 2010, 2016; Gilman 2013; Cregg and Ellison 2018). However, some root shaving methods can stress the plant, resulting in a short-term decline in plant growth (Wilson 1988; Hamilton 1989). In the glasshouse experiment, we found that root shaving reduced the growth of E. reticulatus (biomass and plant height), F. australis (biomass), L. confertus (biomass and plant height) and S. glomulifera (plant height and stem diameter) after 16 weeks. Similarly, in the field experiment, root shaving reduced the growth of E. reticulatus (stem diameter) and S. glomulifera (plant height and stem diameter) after 16 weeks. These reductions in shoot growth after root shaving may be due to the plants redirecting photoassimilates away from shoot growth towards root growth to restore their R:S (Wilson 1988; Hamilton 1989). This is supported by the fact that all the study species with the exception of H. pedula were rapidly able to restore their R:S after root shaving. Further, previous studies have shown that root growth is stimulated after root shaving in urban plant species (Abod and Webster 1990; Blanusa et al. 2007; Cregg and Ellison 2018). Restoring a balanced R:S is critical for plants because it enables them to extract the water and nutrient resources from the soil that are required to maintain their shoot biomass (Struve 2009). Without this allocation shift, the plants would likely have exhibited signs of water stress and/or nutrient deficiency, which were not observed in our experiments. In the case of H. pendula, although it was unable to restore its R:S after root shaving, it did not experience reductions in shoot growth, which suggests it was able to maintain its shoot biomass without needing to compensate for the root loss through greater root growth.

The study species’ ability to restore their R:S after root shaving may have been due to not only their carbohydrate reserves but also their ability to maintain their photosynthetic capacity (Hamilton 1989). This is evident from the fact that the assimilation rate of E. reticulatus, F. macrocarpa, H. pendula and L. confertus were at the levels of the unshaved plants four weeks after treatment. Previous studies have also reported that root shaving does not significantly affect the assimilation rate of urban plant species (Blanusa et al. 2007; Cregg and Ellison 2018; Rouse and Cregg 2021). Further, a plant's ability to maintain its photosynthetic capacity after root pruning, in general, is often aided by having an ample supply of water and nutrients (Richardson-Calfee et al. 2007; Fang et al. 2010; Yin et al. 2014; Adu-Yeboah et al. 2016; Dong et al. 2016), which was the case in our study. As a result of maintaining their photosynthetic capacity after root shaving, the field experiment plants were able in the long-term (i.e. 52 weeks) to compensate for the initial reductions in shoot growth they experienced. Similarly, the shoot growth of the field experiment plants did not differ between the shaving treatments after 120 weeks. Previous studies have reported similar findings, with root shaving not impacting the shoot growth of urban plant species in the longer term (Gilman et al. 2010; Gilman 2013; Cregg and Ellison 2018). These results show that root pruning stress is not universal but rather dependent on the pruning method. That is, unlike harsher pruning methods (e.g. trenching, bare rooting), there is very limited evidence that root shaving causes plant stress (see Andersen et al. 2000).

The effect of biostimulants on urban plant species performance is mixed, with both positive (Ferrini and Nicese 2002; Fraser and Percival 2003) and no effects (Fraser and Percival 2003; Abbey and Rathier 2005; Percival 2006; Banks and Percival 2012, 2014) being reported. The only evidence we found of a biostimulant effect on our study species was that the humic substance biostimulant increased the stem diameter growth of E. reticulatus compared to the protein hydrolysate biostimulant 52 weeks after treatment. Interestingly, there is a large body of literature on edible and ornamental plant species that have reported biostimulants improve plant growth and stress resilience (Calvo et al. 2014; Rose et al. 2014; Colla et al. 2017; Du Jardin 2015; De Pascale et al. 2018; Parađiković et al. 2019). There are several factors (e.g. environmental and soil conditions, method, dosage and frequency of application) that influence the effectiveness of biostimulants on plants (Li et al. 2022). However, the discrepancy in findings between urban and agricultural plant species is most likely due to biostimulant effectiveness being growth form-specific. That is, biostimulants are more likely to have a positive effect on fast-growing plant species than slow-growing woody species (Li et al. 2022).

In this study, we showed that the growth of urban plant species is not impacted by root shaving in the long-term. We also showed that biostimulant application does not affect the performance of urban plant species. This result highlights a significant issue when determining whether biostimulants are beneficial or not. That is, unlike root pruning, the effects of biostimulants on plant performance vary widely due to their interactions with environmental factors (e.g. weather, soil abiotic properties) and the diversity of products that have different modes of action (Ricci et al. 2019). The context-dependent nature of biostimulant effectiveness is reflected in the fact that multiple recent reviews have questioned the standard proof of efficacy used by manufacturers (Yakhin et al. 2017; Van Oosten et al. 2017). Therefore, standardising how we measure the effectiveness of biostimulants to distinguish their true value should be made a priority. Ricci et al. (2019) have proposed a framework that outlines how this may be achieved and will hopefully enable us to better understand the complex array of plant responses to biostimulants. However, at this stage, it remains unlikely that biostimulant application will be a beneficial practice to implement during nursery production and urban greening projects.