Coding regions of HAP-type phytase gene paPhyC from Pantoea agglomerans (ABD85282.1) (Greiner, 2004) and BPP-type phytase gene 168phyA from Bacillus subtilis (CAB13871.1) (Tye et al., 2002) were codon-optimized for expression in A. thaliana using the Codon Adaptation Tool software¹ (Grote et al., 2005). paPhyC and 168phyA coding regions were chemically synthesized (GenScript United States Inc.) as in-frame 5′ fusions with the carrot extensin leader sequence (Richardson et al., 2001) for efficient protein secretion and 3′ fusions with 6x-His (CATCATCATCATCATCAT) and Strep-tag II (TGGTCTCATCCTCAATTTGAAAAG) sequences (Supplementary Figures S1, S2). BamHI and SpeI recognition sites were added on 5′ and 3′ ends of the synthetic constructs, respectively, to facilitate further cloning. Both constructs were placed under the control of the full-length Pht1;2 promoter (-2000 to -1 bp relative to the start codon of AtPht1;2 gene) (Mudge et al., 2003), which was amplified from A. thaliana genomic DNA using primers 5′-CTGCAGGATCACTATACAACTCTGCACT-3′ and 5′-GGATCCCTAAGCCTCTCTTGTCTTTCC-3′ (PstI and BamHI restriction sites are underlined). Both Pht1;2::phytase constructs were inserted into pCBK05 binary vector (Riha et al., 2002) by replacing the vector-specific CaMV35S promoter using PstI and SpeI restriction enzymes, and the derived vector was designated pCEV03 (Supplementary Figure S3). Empty vector containing only the full-length Pht1;2 promoter (without the phytase constructs) was used as a negative “promoter only” control.

All three pCEV03 constructs were introduced into the Agrobacterium tumefaciens GV3101 strain, which was used to transform wild type Arabidopsis thaliana plants (Columbia ecotype) by the modified in planta method (Bechtold and Pelletier, 1998). A. thaliana seeds were cold treated overnight at 4°C, placed in an environmental growth chamber (Panasonic MLR-352, Japan) and grown under a 16-h light/8-h dark photoperiod at 20°C. Primary transformants T1 were selected on 0.5 Murashige and Skoog basal medium supplemented with 25 mg/l of phosphinothricine (BASTA) (Crescent Chemical, United States). BASTA-resistant plant lines were grown for at least three generations (T1, T2, T3) to identify homozygous lines with a single T-DNA insertion site per genome, as judged by a 3:1 segregation ratio for BASTA resistance in T2 plants. Homozygous for the T-DNA insertions plant lines were established from T3 generation and used for further analysis. Reverse transcription PCR (RT-PCR) and Western blotting with His-tag antibody were used to verify bacterial phytase expression in the transgenic lines (see below).

The ability of wild type and transgenic plants to utilize different sources of phosphorus was first determined by growth in hydroponic system (Tocquin et al., 2003). Plants were grown for 25 days in 1 l round sterile containers with liquid “standard” nutrient solution (Tocquin et al., 2003) at pH 5.7. Plants were also supplied with different sources of phosphorus in the medium: Pi (800 μM Na₂HPO₄), phytate (133 μM Na-IHP, Sigma–Aldrich) or grown with no added phosphorus source (No-P control). At least three biological replicates (7–12 plants each) per treatment per plant line were analyzed for various morphological and biochemical parameters.

For growth on perlite, seeds were germinated on sterile plates with 0.5 MS agar in a plant growth chamber (20°C day/20°C night, 16/8 h light/dark conditions) and grown until 4-leaf stage. Plants were then transferred to 6 pots (6 × 6.75 × 6 cm, W/L/H, 3 plants per pot) filled with 100 g of Perlite (Agroperlit, Vita Line) pre-fertilized with liquid “standard” nutrient solution (Tocquin et al., 2003) pH 5.0 ± 0.5 containing sodium phytate (133 μM Na-IHP, Sigma–Aldrich) as the only source of phosphorus. Plants were grown for 21 days in 20°C, 16/8 h light/dark conditions with regular fertilization by phytate-containing “standard” nutrient solution 2–3 times a week. At the end of the experiment, three to six independent replicates (the average of 3 individual plants in each pot) were collected for wild type and every transgenic plant line and analyzed for growth, morphology parameters and phosphorus content.

RNA was extracted from roots grown in hydroponic conditions in the presence of phytate using the TRI reagent solution (Sigma). First strand cDNA synthesis from 1 μg of total RNA was conducted using RevertAid H Minus First Strand cDNA Synthesis Kit (Thermo Fisher Scientific) and Oligo-dT primer. The gene-specific PCR step was performed using the forward primer 5′-ATCACAACCACCACCTCCCTC-3′ specific to the first exon of both constructs (located inside the 2 kb Pht1;2 promoter region) and reverse primers 5′-TCCATAGCCTTCAAAGCTTG-3′ or 5′-CCTTATCTCCATCAATAGCA-3′ specific to the codon-optimized sequences of paPhyC and 168phyA, respectively.

Expression of 168phyA and PaPhyC phytase proteins in plants was confirmed by Western blot analysis. Plants were grown in hydroponic nutrient solution supplied with phytate for 28 days. Plant roots were collected, ground in liquid N₂ with mortar and pestle and transferred to Protein Extraction Buffer (15 mM MES/Ca buffer containing 1 mM cysteine and 1 mM EDTA). Homogenized tissue was then spun at 12000 g for 10 min at 4°C. Supernatant was collected as soluble protein extract and stored in -20°C. Pellet was resuspended in cell wall extraction buffer [15 mM MES (pH 8.5), 1 mM EDTA, 100 mM NaCl, 1% Triton X-100, 0.5 mM CaCl2, 1 mM PMSF, 5 mM cysteine] and spun at 15,000 g for 10 min at 4°C. Supernatant was collected as cell wall protein extract fraction and stored at -20°C. For Western blotting 30 μg of each protein extract was incubated for 5 min at 85°C and subjected to SDS-PAGE (12.5% acrylamide gel). Separated proteins were transferred onto a PVDF membrane for 50 min at 90 V using Mini Trans-Blot Electrophoretic Transfer Cell (Bio-Rad). Membrane was blocked with 5% Skim Milk (Sigma–Aldrich) in PBS-T buffer for 2 h at RT with shaking. PVDF membrane was then incubated with primary antibodies (6x-His Epitope Tag Monoclonal Antibody HIS.H8, Thermo Fisher Scientific) at 1:3,000 dilution for 1 h at RT with shaking. The membrane was washed for 10 min three times in PBS-T buffer and incubated with secondary antibodies [Pierce Goat Anti-Mouse IgG, (H + L), Peroxidase conjugated, Thermo Fisher Scientific] at 1:10,000 dilution for 30 min. After washing in PBS-T and PBS three and two times, respectively, the membrane was visualized using a chromogenic substrate containing SuperSignal West Pico Stable Peroxidase Solution and SuperSignal West Pico Luminol/Enhancer Solution (Thermo Fisher Scientific).

Protein concentration was calculated with a Dc Protein Assay (Bio-Rad) using bovine serum albumin as a standard. Phytase activity was assayed by measuring the amount of released inorganic phosphate (Pi) with a modified ammonium molybdate method (Jog et al., 2005). Briefly, 50 μl of enzyme solution was added to 200 μl of 15 mM MES buffer pH 5.5 with 1.25 mM Na-IHP, 0.5 mM CaCl₂ and incubated at 37°C for 1 h. The reaction was stopped by the addition of 50 μl of 50% TCA. Pellet was spun down for 5 min at 15,000 rpm. Blank controls were prepared by adding 50% TCA prior to the addition of enzyme. Colorimetric reactions with ammonium molybdate solution were then performed as previously described (Jog et al., 2005). Optical density was measured at 820 nm on a model 2550 Microplate Reader (Bio-Rad, United States). A calibration curve was built using concentrations of inorganic phosphate in the range of 5.625–90 μM. One unit (U) of phytase activity was defined as the amount of enzyme necessary to produce 1 μmol of inorganic phosphate per min at 37°C.

For dry weight evaluation shoots of each individual plant were collected separately and washed with dH₂O. Excess water was removed by lightly pressing plants on filter paper. Samples were dried in dry heat oven at 70°C for 48 h and dry weight of each shoot sample was measured on analytical balances. Phosphorus content in plant tissues was estimated by ashing dried plant material at 420°C for 4 h to overnight in a muffle furnace. Ash was then dissolved in 0.9 M H₂SO₄ (w/v, 10 mg of dried tissue/ 1 ml of 0.9 M H₂SO₄) for 24 h in RT. An aliquot of the supernatant was carefully taken and the total phosphorus content was measured by reaction with ammonium molybdate as described for phytase activity assay (Jog et al., 2005). Absorbance at 820 nm was measured and the inorganic phosphate concentration was determined from a calibration curve using KH₂PO₄ as the standard.

Each individual plant was harvested, washed with water and slightly dried with filter paper. Rosettes were gently separated. Plant material was placed on gel documentation system tray and straightened by tweezers. Shoot images were captured using E-box VX2 gel documentation system (VilberLourmat, Germany) with 1.4 MP Sony camera. After taking pictures, morphological parameters of plants (rosette diameter, total area of rosette leaves) were calculated using ImageJ software². Rosette diameter was measured as the mean maximum distance between distal ends of two oppositely positioned rosette leaves. Calculated areas of all rosette leaves in a plant were added up to obtain values for total rosette leaf area. For seed germination analysis several hundred seeds from each line were germinated on sterile plates with 0.5 MS agar for 12 days in a growth chamber (22°C day/20°C night, 14 h light). Only seedlings that formed true first leaves were counted as germinated seeds.

Statistical significance for hydroponics-grown plants was determined using two-way ANOVA with Bonferroni’s correction for multiple comparisons. Data from perlite-grown plants were analyzed by one-way ANOVA with Bonferroni’s correction for multiple comparisons. Significance was set at p < 0.05. All analyses were performed on 3–6 biological replicates using GraphPad Prism version 7.04. Fold change estimates were obtained by dividing the values for transgenic plants by the average of all control A-line values for the same experimental condition.

To test the effects of HAP and BPP phytase expression in A. thaliana on plant morphology and physiology, the coding regions of phytase genes paPhyC from Pantoea agglomerans and 168phyA from Bacillus subtilis were codon-optimized for expression in plants (Chuluuntsetseg et al., 2015; Valeeva et al., 2015), fused with the carrot extensin leader sequence (Supplementary Figures S1, S2) and placed under the control of the full 2 kb inducible root-specific Pht1;2 promoter from A. thaliana (Mudge et al., 2002, 2003). A. thaliana wild type plants (Columbia ecotype) were transformed with T-DNA containing these constructs (designated K-line for 168phyA and G-line for paPhyC) (Supplementary Figure S3) and with a control construct harboring the full Pht1;2 promoter but no phytase gene (designated A-line). Transformed lines harboring a single T-DNA insertion site per genome were identified by analyzing segregation ratios of BASTA resistance in T2 and T3 generations. Phytase mRNA expression was detected by RT-PCR (Supplementary Figure S4A) (Nyamsuren et al., 2015), and phytase protein expression was confirmed by Western blotting (Supplementary Figures S4B,C). Confirmed transgenic lines and control A-lines (harboring just the Pht1;2 promoter) were selected for further morphological and physiological analyses.

We first determined the levels of soluble and cell wall-associated phytase activity in roots of transgenic and control (wild type and A-lines) Arabidopsis plants. All plants were grown in hydroponic conditions with inorganic phosphate (Pi) or phytate (IHP) as the sole source of P for 25 days. Plant growth in the absence of any source of phosphorus in the medium (No-P control) was also attempted, but resulted in immediate developmental abnormalities and plant death by day 10, obstructing further long-term analysis of plants in this condition.

To test the impact of bacterial phytase expression on plant morphology and growth characteristics, we evaluated wild type and transgenic shoots grown in hydroponics for the following parameters: total rosette leaf area and rosette diameter, shoot dry weight and overall phosphorus content.

Wild type A. thaliana plants are largely unable to utilize phosphorus from phytate, and their overall inorganic phosphorus content decreases when plants are grown on phytate as the sole source of phosphorus as compared to Pi (Richardson et al., 2001). To test if transgenic plants expressing 168phyA and PaPhyC phytases are indeed scavenging more phosphorus from phytate than control plants, we measured internal phosphorus content in transgenic and control shoots. When all transgenic and control lines were grown in the presence of Pi, we observed no statistically significant difference in total Pi accumulation in the shoots, which varied from 4.38 to 5.07 μg/plant (Table 1). As expected from previous reports (Richardson et al., 2001; George et al., 2004), total Pi content in wild type and control A-lines grown on phytate decreased substantially (Table 1). While total Pi content in transgenic K- and G-lines grown on phytate also decreased, most transgenic plants expressing high levels of bacterial phytases maintained higher Pi levels as compared to control A-lines. Specifically, phosphorus content in transgenic plants increased over A-lines by 1.90-fold (K1725 line), 1.65-fold (K1115 line), 3.80-fold (G214 line), and 1.71-fold (line G2191) (Table 1). Phosphorus content in transgenic plants with lower levels of phytase activity (K1151 and G251) did not significantly differ from control plants. Overall, these data indicate that, when grown in hydroponic conditions, transgenic plants with high levels of recombinant PaPhyC or 168phyA phytase activity not only display improved growth in the medium with phytate, but do so by accumulating higher net phosphorus content than wild type or corresponding control plants.

To test if expression of bacterial BPP and HAP phytases results in improved plant growth in soil-like conditions, we grew all experimental and control plants in pots containing soilless perlite medium. Unlike other soil-like compounds (i.e., vermiculate), perlite does not release its own nutrients and thus offers a superior choice for manipulating nutrient composition in the growth medium while simultaneously serving as an excellent solid substrate for root development. Hence, perlite is often used as a potting medium in experiments evaluating the effects of phosphorus uptake on plant development (Cabello et al., 2005; Garces-Ruiz et al., 2017). Plants were grown on perlite fertilized with “standard” nutrient solution at pH 5.0 ± 0.5, as higher pH values at 5.7 and above apparently resulted in the formation of insoluble phytate-containing precipitates, as previously reported for sand-vermiculate mixtures (Hayes et al., 2000). After 21 days of growth all control and transgenic plants were evaluated for shoot dry weight, rosette leaf diameter, total leaf area and total phosphorus content. Shoot dry weight of wild type and control plants of A-lines grown on phytate remained within 2.60–6.29 mg/plant range (Table 2). Overall, 3 K- and G-lines grown in perlite on phytate showed significant improvement in dry weight accumulation over wild type or A-line controls (Table 2). Specifically, K1725, K1115 and G2191 lines showed statistically significant increase in shoot dry weight (2.30-, 3.87-, and 3.29-fold increase, respectively), while the other 3 transgenic lines did not substantially differ from wild type and A-line controls. Importantly, the better-performing transgenic lines K1725, K1115 and G2191 are also characterized by the highest levels of transgenic phytase activity when grown on phytate (Figure 1B). This observation further highlights the apparent functional link between efficient phytase expression and growth improvement in conditions when phytate is the only source of phosphorus in the medium.

We next looked at the rosette diameter and leaf area parameters (Table 2). As expected, rosette diameter and leaf area of wild type and control plants grown on phytate did not show substantial variation and ranged between 1.89–2.72 cm and 1.40–1.93 cm², respectively. Among all 6 168phyA and PaPhyC expressing lines, only the two lines with the highest levels of secreted recombinant phytase activity (K1115, G2191) displayed statistically significant increase in rosette diameter and leaf area over values for wild type and control plants.

Finally, we measured total phosphate content in all plants grown on perlite. Total phosphate content of wild type and control A-line plants ranged between 0.28 and 0.51 μg/plant (Table 2). In contrast, plants of all K-lines harbored significantly higher levels of phosphate (1.54 to 2.49-fold increase over A-lines). In addition, plants of G2191 line also displayed a 2.49-fold enrichment in phosphate levels over control values. Altogether, our data indicate that although a substantial degree of variation in growth parameters is observed in transgenic plants grown in hydroponics and soil-like conditions, transgenic lines showing the highest phytase activity levels have the potential to outperform wild type and control plants in terms of biomass production and total plant phosphate content in all conditions tested.

Previous reports have indicated that expression of bacterial phytase genes in plants may cause problems with seed viability (Yip et al., 2003; Lung et al., 2005). To check for any negative effects of 168phyA and PaPhyC expression on seed germination, seeds from control and transgenic lines with the highest levels of phytase activity were planted on 0.5 MS agar medium in Petri dishes and scored for successful germination. Germination rates varied between 97.77% (wild type) and 100% (G214) (Supplementary Table S1). Thus, we conclude that expression of either bacterial phytase in A. thaliana does not lead to any substantial negative impact on seed germination. Overall, our data indicate that expression of both 168phyA and PaPhyC phytases in plants results in higher soluble and cell wall associated phytase activity, improved shoot growth and elevated phosphorus uptake when plants are grown on phytate without negatively impacting seed germination.

Commercial use of microbial phytases is being developed as a promising strategy aimed at increasing crop yield and controlling soil and water pollution. Indeed, much effort has been made to improve phosphorus plant nutrition by co-cultivating plants with bacterial producers of secreted phytases (so-called bio-fertilizers) or by direct addition of purified microbial phytases to soil (Hayes et al., 2000; Zhengao and Huayong, 2001; Idriss et al., 2002; Vessey, 2003; Fuentes-Ramírez and Caballero-Mellado, 2005). While in theory highly promising, this approach has a limited potential due to high costs of recombinant enzyme production and the need to optimize the intricate interactions between plants, bacteria and different soil types (Zimmermann et al., 2003; George et al., 2004). One potentially promising alternative to bio-fertilizers is the use of genetically engineered plants secreting phytases of microbial origin into rhizosphere. Indeed, many recent reports indicate that various plants expressing microbial phytases can efficiently utilize phytate when grown on synthetic media in laboratory conditions. Most of such phytases used in transgenic research are members of the HAP type of phytases (Pen et al., 1993; Verwoerd et al., 1995; Brinch-Pedersen et al., 2000; Ponstein et al., 2002; Hong et al., 2004; Bilyeu et al., 2008; Wang et al., 2013). In addition, a few studies reported successful use of BPP phytases (Yip et al., 2003; Lung et al., 2005; Chan et al., 2006; Belgaroui et al., 2014; Belgaroui et al., 2016) and PAP enzymes (Xiao et al., 2005; Ma et al., 2009, 2012). TTP phytases, which represent a relatively rare family of phytases found mostly in ruminant bacteria (Yanke et al., 1998), have also been utilized (Hong et al., 2004). Thus, although most phytase families have been used in plant biotechnology, it is currently still unclear which phytase type offers the most benefits to transgenic plants while simultaneously causing the fewest side effects to plant physiology and cellular metabolism.

To answer this question, expression of phytases from different enzyme families needs to be analyzed in parallel and under the exact same conditions in plants. Surprisingly, very few studies analyzed simultaneous expression of more than one phytase in planta. In two studies, expression of two plant-derived MtPHY1 and MtPAP1 genes (both purple acid phosphatases) in white clover and in alfalfa resulted in improved phosphorus acquisition and increased biomass when plants where supplied with organic phosphorus (Ma et al., 2009, 2012). Similarly, two HAP phytase genes (phyA from Aspergillus niger and appA from Escherichia coli) were overexpressed under identical conditions in Brassica napus and both resulted in improved soil organic phosphorus utilization (Wang et al., 2013). Finally, in one study expression of two different families of phytases (a cysteine acid phytase SrPf6 from ruminant bacterium Selenomonas ruminantium and a HAP phytase appA from E. coli) in rice seeds resulted in no developmental or seed abnormalities, though growth on organic phosphorus was not evaluated (Hong et al., 2004). However, to the best of our knowledge, comparative in planta analysis of expression phenotypes of members of the two most biotechnologically important families of phytases, HAP and BPP, has previously not been described.

In this work we have directly compared expression in plants of a HAP phytase from Pantoea agglomerans and a BPP phytase from Bacillus subtilis using the same promoter and identical growth conditions. Specifically, we tested whether expression of either bacterial phytase in A. thaliana leads to not only improved plant growth on phytate but also results in better inorganic phosphorus accumulation in the shoots. Because expression level of microbial proteins in plants may not be optimal owing to different codon usage preferences in plants and bacteria (Gouy and Gauter, 1982; Hamada et al., 2005), we have performed codon optimization of coding regions for both genes without changing their amino acid sequences prior to the introduction of transgenes into plant genomes. The choice of promoter represents one of the most important ways to optimize heterologous gene expression. While many root-specific promoters have been used in different studies to direct phytase expression (Mudge et al., 2003; Zimmermann et al., 2003; Xiao et al., 2005; Li et al., 2009), the inducible Pht1;2 promoter is not only one of the most well-characterized, but has also been shown to respond very well to conditions of phosphorus starvation in A. thaliana (Mudge et al., 2002, 2003). Thus, to specifically express microbial phytases in plant roots, both paPhyC and 168PhyA genes have been cloned under the control of the full 2 kb A. thaliana promoter Pht1;2 (Mudge et al., 2003). To facilitate secretion of transgenic proteins into the rhizosphere, we fused the coding region of both phytases with carrot extensin leader sequence (Chen and Varner, 1985). To better characterize the influence of transgenic phytase expression on plant morphology and physiology, we turned to growing transgenic and control plants on hydroponics. This experimental set-up has allowed us to obtain large volume of plant root systems for protein expression analysis, as well as for morphological and biochemical assays. In parallel, we have evaluated transgenic plant performance in soilless perlite medium, which mimics more natural plant growth conditions and whose nutrient composition can be easily manipulated.

Our data indicate that transgenic plants of K1725, K1115, and K1151 lines (expressing 168phyA phytase) and of G214 and G2191 lines (expressing PaPhyC phytase) display much higher phytase activity levels in root extracts (especially in cell-wall bound fractions) on phytate medium as compared to wild type and empty vector control plants. These results further confirm that expression of both bacterial phytases, as expected, was not only induced under phosphate-starvation conditions but resulted in protein secretion across plasma membrane. In contrast, phytase activity in G251 line was not significantly induced, possibly highlighting important differences in specific T-DNA integration sites. Overall, our data correlate well with previously published analysis of transgenic A. thaliana plants expressing a HAP-type Aspergillus niger phytase under the control of the same Pht1;2 promoter, which similarly had much higher activity on low Pi medium (Mudge et al., 2003).

Expression of bacterial phytases in plants has previously been associated with important changes in plant morphology, physiology and phosphorus content. For instance, expression of 168phyA phytase in tobacco and in A. thaliana resulted in increased shoot biomass (up to 1.7–2.2 fold) and 27–36% higher phosphorus content as compared to wild type (Yip et al., 2003; Lung et al., 2005). Here we show that expression of bacterial genes encoding a HAP phytase from P. agglomerans and a BPP-type phytase from B. subtilis resulted in improved plant growth on hydroponic medium with phytate as the sole source of phosphorus. Specifically, we observed increased rosette diameter (2.18–2.79 fold) and leaf area (2.50–5.10 fold), increased dry weight (1.86–3.35 fold) and significantly elevated shoot phosphorus content (1.65–1.90 fold), as compared to control plants, but only in transgenic plants with relatively high levels of phytase expression. Similarly, when transgenic plants with high levels of bacterial phytase expression (specifically, lines K1725, K1115, G2191 and for some parameters also lines K1151 and G214) were grown on perlite, we observed comparable levels of growth improvements, including up to 3.87-fold increase in shoot dry weight, up to 1.61-fold increase in rosette diameter, up to 2.79-fold increase in rosette leaf area and up to 2.49-fold increase in total P content. Taken together, our data clearly indicate that plants efficiently secreting both 168phyA and PaPhyC phytases are capable of cleaving phytate into inorganic phosphate and myo-inositol, leading to an increased overall inorganic phosphorus intake and improved growth.

Several lines of evidence support the notion that the presence of both transgenic phytases does not lead to any adverse effects on plant morphology and physiology. First, leaf morphology of all transgenic plants grown on Pi or phytate remained normal. Second, we observed no change in seed germination efficiency of transgenic plants. Third, the majority of phytase activity appears to be cell wall associated, suggesting that heterologous expression of bacterial phytases in these conditions is unlikely to cause any major physiological changes in the cytoplasm. We conclude that expression of bacterial phytases in A. thaliana leads to improved plant growth and phosphorus acquisition on hydroponic medium with phytate without any major detrimental outcomes. Thus, our results provide additional experimental support for the notion that the power of biotechnology can be harnessed to improve plant growth on marginal soils with little Pi but abundant phytate reserves.

While physiological phenotypes of PaPhyC expression in plants have not been evaluated prior to this study, many other members of the HAP family of phytases have been extensively used to engineer both model plants and crops. In most cases, expression of a HAP-type phytase in plants led to important beneficial changes in plant morphology and metabolism, including better P accumulation and growth on phytate-containing medium (Gontia et al., 2012; Yao et al., 2012). 168phyA phytase has previously been expressed in tobacco and A. thaliana, though under different promoters (Yip et al., 2003; Lung et al., 2005; Chan et al., 2006). A closely related phytase from B. subtilis strain 417 (72% amino acid identity and 84% similarity to 168phyA phytase) has also recently been used for this purpose (Belgaroui et al., 2014, 2016). When grown on phytate, plants expressing these BPP-type phytases displayed twofold higher biomass levels (Belgaroui et al., 2016), as well as a significant increase in shoot P concentration compared to control plants (Lung et al., 2005; Belgaroui et al., 2016). However, in some cases these phenotypes were accompanied by negative effects on plant physiology, including smaller size and lower germination rates of transgenic tobacco seeds expressing 168phyA phytase (Yip et al., 2003; Lung et al., 2005). Thus, an important conclusion from our study is that expressing 168phyA phytase as a secreted protein under the control of inducible Pht1;2 promoter does not cause any apparent adverse physiological effects and represents an effective route to utilize a BPP-type enzyme in applications of plant biotechnology.

Our data on 168phyA and PaPhyC expression in A. thaliana indicate that both phytases, when highly expressed under identical conditions and from the same promoter, clearly improved plant growth and P acquisition from phytate to a similar degree, both in hydroponics and perlite. Interestingly, the major difference between 168phyA and PaPhyC phytase expression in our experiments appears to be the phenotype of dry weight accumulation in conditions of hydroponics, when plants highly expressing PaPhyC are characterized by overall significantly higher shoot dry weight than their 168phyA-expressing counterparts. Taken at face value, this observation may suggest that a HAP-type phytase could be more beneficial when increased shoot biomass is the desired outcome of transgenic manipulations. However, when the same plants were grown on perlite with phytate as the only source of phosphorus, we did not detect any significant differences in shoot dry weight accumulation between 168phyA and PaPhyC expressing plants (though both transgenics clearly improved plant growth over control lines). This observation supports and correlates well with previous data indicating that growth improvement in laboratory conditions in vitro does not necessarily translate to growth improvement in more natural field conditions. One possible explanation for the observed differences in dry weight accumulation between 168phyA and PaPhyC expressing plants in hydroponics and perlite is that enzyme’s access to the substrate may be more limited in perlite than in hydroponic conditions. Indeed, in a hydroponic system roots can move more freely than when plants are grown in soil-like conditions, affecting access to the substrate and possibly even its local concentration. This potentially can help explain both our data and results of other research groups when excellent growth phenotypes of many transgenic plants grown in laboratory conditions are often compromised when these same plants are grown in soil (George et al., 2004, 2005), though some of these setbacks can apparently be modulated or even reversed through other considerations, such as a choice of promoter (Wang et al., 2009), phytase gene, signal sequences or soil types (Ma et al., 2012).

It should be noted that additional factors can influence the effects of transgenic phytase expression on plant growth. For example, some bacterial phytases expressed in plants are often post-translationally modified (Chan et al., 2006; Chen et al., 2008), raising the possibility that the observed differences in plant morphology may at least partially arise from distinct protein modification pathways affecting individual phytase molecules. Furthermore, individual enzyme’s preferences for optimal pH and temperature can have a major impact on phytase activity in plants. For example, PaPhyC phytase has a single pH optimum at pH 4.5 and temperature optimum around 60°C when purified from bacteria (Greiner, 2004). On the other hand, 168phyA phytase expressed in Bacillus preferred more neutral pH and similarly high optimal temperature (Tye et al., 2002). As pH of the hydroponic growth medium at the beginning of our experiments was set at 5.7, and pH of perlite medium was set even lower, these environmental parameters could have differentially affected recombinant PaPhyC and 168phyA enzymes, resulting in different outcomes. Finally, as tolerance to organic acids (specifically, citrate and malate) secreted by roots under phosphorus stress appears to be a determining factor affecting performance of at least some phytase enzymes in the rhizosphere (Tang et al., 2006; Lei et al., 2013), the ability of 168phyA and PaPhyC to tolerate these compounds can also have important implications. Nevertheless, our data provide the first glimpse into physiological differences between expressing individual members of HAP and BPP phytase families in plants and can serve as a useful guide into future research in this area.

Overall, while various biochemical constraints are clearly important and need to be taken into consideration when designing future plant biotechnologies, our data clearly demonstrate that for both phytases those transgenic lines that harbor the highest levels of recombinant phytase activity also consistently demonstrate greater growth improvement in both hydroponics and soil-like conditions. Thus, our data strongly reinforce the notion that the actual amount of enzyme activity is essential and can be a limiting factor in some transgenic studies. It is likely that while in hydroponic conditions the transgenic effects are more evident at a wider range of transgenic phytase activity levels, in a more real soil-like situations the amount of available active phytase needs to pass a certain threshold level to have a more significant impact on plant physiology. In general, while it is not uncommon for transgenic plant lines to display both different levels of phytase expression and different effects on plant growth in P-deficient conditions (George et al., 2004), our findings provide valuable quantitative comparative data on important correlations between higher phytase expression levels and greater improvements in growth phenotypes when plants are grown on phytate.