3.1. Physiological Parameters
After flooding, the plants oxygen concentrations in the water were measured continuously because, as with waterlogging stress in nature, no adjustments were done to change oxygen concentration in one direction or another. After four hours of waterlogging, oxygen concentration was 84%, after 28 h it decreased to 22% and on the last measurement point, after 52 h, it was down to 2%. Typical symptoms of flooding stress were found for maize after three days of waterlogging. The roots of flooded plants tended to become negatively gravitropic (
Figure 1A). Shoot length was compared for control and flooded plants at four, 28 and 52 h. The ratio of the average shoot growth (4 h: 1.0, 28 h: 1.0 and 52 h: 1.1) of control
versus submerged plants did not change significantly (
Figure 1B), but showed already a tendency of a decreased growth of the stressed plants. Furthermore, after 52 h of flooding stress, the shoot stem diameter was increased about 24% within stressed plants in comparison to the control plants (
Figure 1C,D). Additional to these parameters, chlorophyll a and chlorophyll b content was determined by spectrophotometric measurements (
Figure 1E). At the first day of flooding stress, the chlorophyll b concentration decreased about 17.2% after four hours, if compared to chlorophyll a. At the second day, chlorophyll b content was decreased about 24.3% compared to chlorophyll a. At the third day, chlorophyll b content was lowered about 17.8% in comparison to the chlorophyll a content.
Figure 1.
Morphological adaption of maize to submergence. (a) Phenotype of control plants and plants stressed by submergence at the end of the growing period; (b) Shoot length was measured for control and stressed plants over the three days of flooding; (c) Shoot basis of control and submerged plants; (d) Comparison of basal shoots of control and stressed samples at the end of the experiment; (e) Chlorophyll a/b ratio. All measurements were done for n ≥ 20 biological replicates. Except for (e) measurements were done for n ≥ 5 biological replicates. Control, light grey, flooded plants, dark grey. Significant changes were marked with an asterisk.
Figure 1.
Morphological adaption of maize to submergence. (a) Phenotype of control plants and plants stressed by submergence at the end of the growing period; (b) Shoot length was measured for control and stressed plants over the three days of flooding; (c) Shoot basis of control and submerged plants; (d) Comparison of basal shoots of control and stressed samples at the end of the experiment; (e) Chlorophyll a/b ratio. All measurements were done for n ≥ 20 biological replicates. Except for (e) measurements were done for n ≥ 5 biological replicates. Control, light grey, flooded plants, dark grey. Significant changes were marked with an asterisk.
Maize plants showed typical phenotypes after three days of waterlogging (
Figure 1). Growth of the shoots was compared for control and flooded plants for 4, 28 and 52 h (three days of flooding). The ratio of the average shoot growth of stressed and control plants did not change significantly. Shoot growth was only slightly decreased for water logged plants after 52 h (
Figure 1B). These observations confirm published data for water logged maize [
27]. Reduced elongation growth is based on the negative effect of flooding on photosynthesis and is in accordance with the low oxygen quiescence syndrome of maize [
9,
10,
27].
The decrease of chlorophyll a/b ratio (
Figure 2) was shown to be a typical reaction to flooding stress in the past [
6,
27]. The decrease of the chlorophyll a/b ratio seems to be a good marker as its change appears shortly after the plant is exposed to flooding stress, but validations are usually needed, because of variation in reaction to flooding stress in different species. Also, the thickening of the shoot stem diameter after a few days of flooding is in accordance with published data for maize [
27]. The thickening of the basal shoot was shown to be based on the aerenchyma formation in the root cortex, which is the most studied morphological response to flooding stress [
27,
34]. Aerenchyma provides a continuous system of interconnected aerial spaces with a lower resistance for oxygen transport. Aerenchyma formation allows root growth and soil exploration under anaerobic conditions by oxygen transport from aerial shoots to submerged roots [
27,
34].
In the past, it was shown that continuous flooding over time causes a decrease in photosynthetic capacity of mesophyll cells and finally to an overall reduction of photosynthetic activity [
27]. The lower photosynthetic activity is based on the lower chlorophyll content, reduced activity of carboxylation enzymes and oxidative damage of photosystem II by ROS. Low photon utilisation of flooded plants results mostly in ROS production [
35]. The level of ROS, like superoxide anion radical, singlet oxygen, hydrogen peroxide and hydroxyl radical, which are highly reactive and provoke damage to various molecules, is tightly managed to protect the cells against oxidative stress. Plants contain antioxidants like ascorbate, glutathione and membrane embedded quinones (e.g., tocopherols and ubiquinone) and enzymes with ability to scavenge ROS and regenerate the antioxidants [
28,
30]. Peroxidases are part of this ROS scavenging mechanism, but they also play a role in cell wall loosening and reorganisation, such as needed for the formation of aerenchyma.
3.2. Differential Regulation of Soluble Peroxidases—1D and 2D PAGE Analysis
Peroxidases play roles in the ROS scavenging mechanism and in cell wall loosening and reorganisation. Data at hand showed alterations of peroxidase profiles, increases in abundance of specific peroxidases and increase or decrease of guaiacol peroxidase activities under waterlogging conditions. These observations were in accordance to published data [
36,
37].
Molecular mass of peroxidases and over all profiles were observed for control and stressed plants for all three time points. Four guaiacol peroxidase bands were detected in all samples after separation by modified SDS-PAGE (
Figure 2A; band A, B, D, E). The peroxidase profiles were different for the observed time points (
Figure 2A). From the relative stress to control (s/c) ratios, relative abundance change was calculated for the bands of the modified SDS-PAGE (
Figure 2B). The strongest regulated band of peroxidase abundance was band B, with 133 kDa (
Figure 2B). This band was significantly decreased after four hours of waterlogging, whereas it was increased after 28 and 52 h of waterlogging. Furthermore, peroxidase profile of four hours waterlogging exhibited a decreased band C, independent of the flooding stress. Overall, bands A–D were significantly decreased in number after four hours of waterlogging (
Figure 2B). After 28 h of waterlogging, stress intensity of bands B–D were significantly increased. At 52 h of waterlogging, only band B was significantly changed. Band E was not significantly changed at any of the observed time points. The overall amount of peroxidase activity per time point, calculated from all bands per lane from all technical/biological replicates, result in the following order: stress day 2 ≥ stress day 3 ≥ control 3 ≥ control 2 ≥ control 1 ≥ stress day 1.
Figure 2.
First dimension gel electrophoresis and guaiacol/H2O2 staining (a) Guaiacol/H2O2 staining after separation by modified SDS-PAGE. The pre-stained marker is shown on the left, indicated with M at the top of the gel. Significantly detected guaiacol bands were amounted with the letters of A–E, referring to their mass; (b) Relative activity of the significantly detected bands A–E in the modified SDS-PAGE (n ≥ 3). The corresponding bands are indicated on the x-axis. Dark grey, s1/c1, light grey, s2/c2, middle grey, s3/c3 (s, stress, c, control, 1–3, day after stress induction); (c) Guaiacol/H2O2 staining after separation by native isoelectric focusing polyacrylamide gel electrophoresis (IEF-PAGE). The picture was inverted to enhance the visibility. Next to the pI, peroxidase identifiers are indicated on the left hand; (d) Relative activity of the significantly detected bands with the pI of 9.6–5.9 in the native IEF (n ≥ 3). The corresponding bands are indicated on the x-axis. Dark grey, s1/c1, light grey, s2/c2, middle grey, s3/c3 (s, stress, c, control, 1, 4 h, 2, 28 h, 3, 52 h). For the gels, the type of sample was indicated at the top with control or stress. At the bottom of the gel, the day after stress induction was specified. Significant changes between control and the associated stressed sample were marked with an asterisk (student’s t-test).
Figure 2.
First dimension gel electrophoresis and guaiacol/H2O2 staining (a) Guaiacol/H2O2 staining after separation by modified SDS-PAGE. The pre-stained marker is shown on the left, indicated with M at the top of the gel. Significantly detected guaiacol bands were amounted with the letters of A–E, referring to their mass; (b) Relative activity of the significantly detected bands A–E in the modified SDS-PAGE (n ≥ 3). The corresponding bands are indicated on the x-axis. Dark grey, s1/c1, light grey, s2/c2, middle grey, s3/c3 (s, stress, c, control, 1–3, day after stress induction); (c) Guaiacol/H2O2 staining after separation by native isoelectric focusing polyacrylamide gel electrophoresis (IEF-PAGE). The picture was inverted to enhance the visibility. Next to the pI, peroxidase identifiers are indicated on the left hand; (d) Relative activity of the significantly detected bands with the pI of 9.6–5.9 in the native IEF (n ≥ 3). The corresponding bands are indicated on the x-axis. Dark grey, s1/c1, light grey, s2/c2, middle grey, s3/c3 (s, stress, c, control, 1, 4 h, 2, 28 h, 3, 52 h). For the gels, the type of sample was indicated at the top with control or stress. At the bottom of the gel, the day after stress induction was specified. Significant changes between control and the associated stressed sample were marked with an asterisk (student’s t-test).

Each lane of modified SDS-PAGE was cut into four pieces, digested and used for identification of proteins by LC-MS (
Supplemental Table S1), but peroxidases were not identified. Possible explanations for the lag of peroxidase identifications are the relatively high sensitivity of the guaiacol staining in comparison to standard staining, e.g., CCB, overlay of high abundant proteins with the same molecular mass and inefficient tryptic digestion based on the nature of the non-reducing SDS-PAGE.
Similar to the modified SDS-PAGE, peroxidase bands of the first dimension native IEF-PAGE were used to obtain isoelectric points (pI) and corresponding peroxidase profiles (
Figure 2C;
Table 1). Overall peroxidase profiles were comparable to the profiles of the modified SDS-PAGE (
Figure 2B). Samples of the first time point showed a different peroxidase profile from the samples of the two following time points, independent of the sample treatment. Semi-quantitative analysis of the activity bands was performed for the native IEF-PAGE, as described for modified SDS-PAGE (
Figure 2C). Significant changes in activity were observed after four hours of waterlogging at pI 9.6 (PrxF1) and 7.0, which were both decreased in the stressed sample. At 28 h of waterlogging, only the band with the pI of 8.0 (PrxF2) was significantly changed, if the ration of stress to control was compared. The band with the pI of 9.2 was only significantly changed at 52 h after induction of waterlogging. The band with the pI of 7.8 (PrxF3) was similar to the band with pI 9.2 significantly increased after 52 h in the waterlogged sample, if compared to the control (
Figure 2D).
Table 1.
Summary of peroxidase properties separated by one-dimensional gel-electrophoresis.
Table 1.
Summary of peroxidase properties separated by one-dimensional gel-electrophoresis.
pIex Native IEF | kDa Modified SDS-PAGE 1D | kDa hrCNE |
---|
9.6 ± 0.3 | 183 ± 7 | 637 ± 7 |
9.2 ± 0.4 | 133 ± 5 | 330 ± 7 |
8.0 ± 0.2 | 68 ± 1 | 431 ± 8 |
7.8 ± 0.2 | 55 ± 1 | 219 ± 8 |
7.0 ± 0.1 | 47 ± 2 | 200 ± 9 |
6.1 ± 0.1 | | 162 ± 8 |
| | 136 ± 0.6 |
| 125 ± 2 |
| 117 ± 1.7 |
| 32 ± 1.7 |
Aside the specific regulations by flooding, peroxidases with acidic pIs showed a regulation independent of flooding on day two (
Figure 2). It is possible that these peroxidases are differentially regulated depending on the developmental stage of the shoot [
14]. Plants were grown in the glass house; therefore, changes in light conditions are also an option for this change, but daily measurements showed light intensity was comparable at all three time points (~1000 µmol/m
2∙s).
In contrast to modified SDS-PAGE that allows estimation of peroxidase abundance, native PAGE allows estimation of peroxidase activities by quantification of the intensity of the guaiacol peroxidase bands [
21]. As shown in
Figure 2, nearly all isoenzymes increased by waterlogged conditions from time point one to three, showing an overall induction of soluble peroxidases. This was observed for both methods, modified SDS-PAGE and native IEF-PAGE, suggesting a relation between peroxidase abundance and activity. Abundance and activity are not related for all proteins, especially if proteins are activated by post-translational modifications [
38].
First dimension hrCNE was used to calculate the native molecular mass of guaiacol peroxidases (
Table 1). Based on the resolution and the high activity of the peroxidases (saturation of the bands), hrCNE could not be used for quantitation. Two different amounts of total protein (25 µg and 40 µg) were loaded to the gels to ensure the detection of both strong and faint bands (strong activity can cover light activity). Finally, 10 bands, ranging from 32–637 kDa, were detected (
Table 1).
Aside the band at 637 kDa, bands of 133 kDa and higher were detected in the modified SDS-PAGE. In both cases, molecular mass detected is fairly high for peroxidases, indicating an association with a protein complex [
39,
40,
41]. This protein band may also present peroxidase aggregates or polymers. These results were confirmed by two dimensional gels, namely native IEF/modified SDS-PAGE and native IEF/hrCNE (
Figure 3,
Table 2). In the native IEF/modified SDS-PAGE combination, two peroxidases with pI 8.0 (PrxF2)/133 kDa and pI 7.8 (PrxF3)/85 kDa were detected, suggesting from their masses to be a dimer and trimer, as the identified peroxidase have an theoretical molecular mass of 27–38 kDa (
Table 3). The same spots were detectable in the hrCNE with a native mass of 200 kDa.
Peroxidase profiles in the second dimension hrCNE varied for the different samples and confirmed the results of the first dimension native IEF-PAGE. The smallest amount of peroxidase spots was detectable in the samples after 4 h independent of the stress, whereas the highest amount of spots was detected after 52 h (
Supplemental Figure S2). The spot with the pI at pH 8.0/7.8 (PrxF2/F3) and at pI 9.2 were clearly separated in the second dimension into two spots (pI 9.2, J, N; pI 8.0, O, Q; pI 7.8, P,R) with different native molecular masses (
Figure 3C,
Table 2). Spots in the second dimension were only used to get a better view on the isoenzymes with similar pI that could not be separated by native IEF-PAGE. Based on gel to gel variation, these gels were not used for quantitation, and identification by MS was not successful.
Table 2.
Summary of peroxidase properties separated by two-dimensional gel-electrophoresis.
Table 2.
Summary of peroxidase properties separated by two-dimensional gel-electrophoresis.
Spot Name | pIex Native IEF | kDa Modified SDS-PAGE | kDa hrCNE |
---|
H | 9.8 ± 0.2 | n.d. | 637 ± 10 |
J | 9.2 ± 0.1 | n.d. | 637 ± 6 |
K | 8.8 ± 0.2 | n.d. | 637 ± 5 |
L | 9.6 ± 0.2 | n.d. | 440 ± 5 |
M | 9.6 ± 0.2 | n.d. | 330 ± 8 |
N | 9.2 ± 0.1 | n.d. | 370 ± 7 |
O | 8.0 ± 0.3 | n.d. | 370 ± 7 |
P | 7.8 ± 0.3 | n.d. | 330 ± 8 |
F SDS−PAGE/Q hrCNE | 8.0 ± 0.3 | 133 ± 8 | 200 ± 4 |
G SDS−PAGE/R hrCNE | 7.8 ± 0.3 | 85 ± 4 | 200 ± 6 |
S | 6.1 ± 0.1 | n.d. | 139 ± 5 |
T | 5.9 ± 0.2 | n.d. | 115 ± 2 |
Figure 3.
Second dimension gel electrophoresis for samples exposed to three days of waterlogged soil. (a) Guaiacol staining of the second dimension modified SDS-PAGE after separation by IEF-PAGE in the first dimension. The pI of the guaiacol detected spots in the first dimension was indicated at the top of the gel; (b) CCB staining of the gel shown in (c); (c) Guaiacol staining of the second dimension hrCNE after separation by IEF-PAGE in the first dimension. The pI of the guaiacol detected spots in the first dimension is indicated at the top of the gel.
Figure 3.
Second dimension gel electrophoresis for samples exposed to three days of waterlogged soil. (a) Guaiacol staining of the second dimension modified SDS-PAGE after separation by IEF-PAGE in the first dimension. The pI of the guaiacol detected spots in the first dimension was indicated at the top of the gel; (b) CCB staining of the gel shown in (c); (c) Guaiacol staining of the second dimension hrCNE after separation by IEF-PAGE in the first dimension. The pI of the guaiacol detected spots in the first dimension is indicated at the top of the gel.
3.4. SEC and Identification of Peroxidases by LC-MS
To confirm the results from the gel electrophoresis, samples were separated by SEC. Peroxidase elution from the column was followed by guaiacol/H
2O
2 micro assay. Analysis of the different samples and biological replicates showed molecular mass from 40–287 kDa with significant variation between the separations. Furthermore, different peroxidase could not be clearly separated (
Supplemental Figures S4 and S5). Additionally, active fractions were separated by one dimensional modified SDS-PAGE, native IEF and hrCNE and peroxidases detected by guaiacol/H
2O
2 in gel staining (
Supplemental Figure S5, Table S2). Observed profiles showed strong similarities independent of the separated sample (control, stress). Molecular mass calculated for the detected bands confirmed bands from one dimensional electrophoresis separation without SEC (
Table 2). Aside that, molecular mass calculated from the fraction number of the SEC varied strongly from the detected bands in the gels. Finally, only bands with a molecular mass above 120 kDa were detected after gel electrophoresis of SEC fractions, independent of the electrophoresis method. Native IEF separation was not possible for most of the samples based on a disturbed electric flow. If separation was possible, activity was detected at pI 6.1, 7.8, 8.0 and 9.6, which confirmed the primary results mentioned above. All spots detected in the native IEF were identified, after tryptic digestion, by MS as peroxidases. Experimental and theoretical properties of the identified peroxidases were summarized in
Table 3, while the complete MS data set for identification of the peroxidases can be found in the Supplemental (
Supplemental Table S3). The pre-separation of SEC overlaying proteins with similar pI, but different molecular mass, meant they were excluded from the separation of native IEF without diminishing the concentration of the protein. Therefore, the chance of identifying a specific protein, e.g., peroxidase, was much higher than in a first dimension modified SDS-PAGE.
In most activity spots, multiple peroxidases were identified (
Table 3). With the experimental pI of the identified peroxidases (PrxF1-F4), they can be assigned to the bands found in the native IEF without pre-separation by SEC. Besides class III peroxidases, also ascorbate peroxidases (APx, class I peroxidases) were identified (ZmAPx01 and 02). These peroxidases play a major role under oxidative stress and have been shown to be regulated under stress conditions [
35,
43]. Even APx was identified in the bands PrxF2 and PrxF3; usually they cannot use guaiacol as substrate. In soybean, flooding stress regulated APx [
22,
23]. We suggest that the APxs identified do not contribute to the detected activity, which would be in accordance with earlier results. Additionally, a plasma membrane associated peroxidase (ZmPrx66) was identified in the analysed soluble fraction, which might be due to (i) a contamination or (ii) the proteins disband under specific conditions from the plasma membrane. If the second point is the case, it will have major influence on the understanding of the stress–peroxidase relation. Aside ZmPrx66, APx1 and APx2, eight more peroxidases were identified in the spots Prx F1 to Prx F4. ZmPrx06 (also named peroxidase J), ZmPrx118, ZmPrx97, ZmPrx124, ZmPrx125, ZmPrx07, ZmPrx38, ZmPrx106 were identified in the maize genome but further information on these soluble peroxidases are not known [
44,
45,
46]. Based on KEGG (Kyoto Encyclopedia of Genes and Genomes) calculations related pathways for ZmPrx118 are the phenylpropanoid [
47] and the lignin biosynthesis [
48], as well as the phenylalanine metabolism [
49]. ZmPrx07 was identified by the NCBI blast as ZmPrx66 precursor and showed 99% similarity to ZmPrx66, making it highly reasonable that the identified peroxidase in the spot Prx F2 is the plasma membrane associated ZmPrx66. ZmPrx42 identified in the band Prx F2 was predicted before as pmPOX3-1 [
42]. In both cases, the functions discussed were removal of H
2O
2, oxidation of toxic reductants, biosynthesis and degradation of lignin, suberisation, auxin catabolism, response to environmental stresses such as wounding, pathogen attack and oxidative stress. These functions might be dependent on each isoenzyme/isoform in each plant tissue. Three of the identified peroxidases have been shown to be induced by biotic or abiotic stress factors (
Table 3). According to the PeroxiBase, ZmPrx97, identified in band PrxF1 with the pI of 9.6, and ZmPrx66, ZmPrx42, identified in the band PrxF2 with the pI of 8.0, are induced by drought and salt stress. Our former data showed alterations of ZmPrx66 abundance at washed plasma membranes by elicitors, salicylic acid and H
2O
2 [
25].
Class III peroxidases may build a complex functional network of different isoenzymes that appears tightly regulated under stress conditions. Depending on a stressor and plant stress responses, distinct isoperoxidases seem to be up-regulated and/or down-regulated. This was shown for maize under submerged conditions (
Figure 2 and
Figure 3), by different signalling compounds and by oxidative stress [
25]. The observed soluble peroxidases have not been able to be assigned to a specific localisation in the cell up to now. Therefore, separation of the peroxidase function in the protective cycle or in the flooding induced leaf growth has to be further investigated. Increased lipid peroxidation and guaiacol peroxidase and APX activity have been demonstrated for maize by flooding in young maize seedlings, but resulting peroxidases were not identified [
50]. Thus, ROS scavenging may be one of the major functions of guaiacol peroxidases induced under waterlogging conditions. Peroxidases may also be involved in the process of adaptation. Aerenchyma formation is correlated to programmed cell death (
i.e., ROS production) and cell wall stiffening. In cell wall fractions of pea (
Pisum sativum L.) roots, alkaline isoperoxidases of ionically bound fraction appeared to be involved in elongation growth, whereas covalently bound peroxidases with acidic pI were suggested to be involved in cell wall related functions [
51]. Furthermore, extracellular isoperoxidases have been demonstrated to be involved in ROS production [
41,
52]. ROS production has been demonstrated during root hair formation [
53]. Thus, functions in formation of adventive roots may also be possible. Localisation and biochemical properties of flood-induced isoperoxidases will need further studies to clarify their physiological functions in maize.
Table 3.
Identified peroxidases by LC-MS. Separated samples by size exclusion chromatography were followed by native IEF and stained with guaiacol/H
2O
2. Detected spots were tryptical digested and analysed by LC-ESI-MS/MS. Identified peroxidases and their properties are listed in the table. Detailed information about the MS results can be found in the
Supplemental Table S3. Name: name used in the publication; pI exp: experimental pI resulting from the calculation of the activity band in native IEF after SEC separation; MW
exp: experimental molecular mass (kDa), resulting from the SEC separation; Accession: accession for the peroxidase in the searched database; pI
th: theoretical pI given by the database; MW
th: theoretical molecular mass (kDa) given by the database; Peptides: amount of identified peptides; Class: classification of the peroxidase identified; Localisation: known cellular localisation; Inducers/Repressors: known inducers or repressors of the identified peroxidase based on information provided by peroxibase [
54].
Table 3.
Identified peroxidases by LC-MS. Separated samples by size exclusion chromatography were followed by native IEF and stained with guaiacol/H2O2. Detected spots were tryptical digested and analysed by LC-ESI-MS/MS. Identified peroxidases and their properties are listed in the table. Detailed information about the MS results can be found in the Supplemental Table S3. Name: name used in the publication; pI exp: experimental pI resulting from the calculation of the activity band in native IEF after SEC separation; MWexp: experimental molecular mass (kDa), resulting from the SEC separation; Accession: accession for the peroxidase in the searched database; pIth: theoretical pI given by the database; MWth: theoretical molecular mass (kDa) given by the database; Peptides: amount of identified peptides; Class: classification of the peroxidase identified; Localisation: known cellular localisation; Inducers/Repressors: known inducers or repressors of the identified peroxidase based on information provided by peroxibase [54].
Band | pI exp | MW exp | Accession | Database | pI th | MW th | Peptides | Class | Localisation | Inducers/Repressors |
---|
Prx F1 | 9.6 | 34–51 | ZmPrx06 | Peroxibase | 6.2 | 33 | 6 | III | - | induced by cyst nematode infection, pathogen interaction |
ZmPrx118 | 5.5 | 37 | 10 | III | - | - |
ZmPrx97 | 6.6 | 38 | 12 | III | - | induced by salt stress |
ZmPrx124 | 4.7 | 37 | 15 | III | - | - |
ZmPrx125 | 8.6 | 34 | 10 | III | - | - |
Prx F2 | 8.0 | 34–58 | ZmPrx66 | UniProt | 8.0 | 33 | 2 | III | PM | induced by drought, elicitors, salicylic acid, wounding and H2O2 |
ZmPrx42 | Peroxibase | 5.3 | 33 | 9 | III | PM | - |
ZmPrx07 | Peroxibase | 8.0 | 34 | 12 | III | - | - |
ZmAPx01 | UniProt | 5.7 | 27 | 27 | I | cytosolic | - |
ZmAPx02 | UniProt | 5.7 | 28 | 9 | I | cytosolic | - |
Prx F3 | 7.8 | 34–58 | ZmAPx01 | UniProt/Peroxibase | 5.7 | 27 | 8 | I | cytosolic | - |
ZmPrx38 | Peroxibase | 9.2 | 38 | 10 | III | - | - |
ZmPrx07 | Peroxibase | 8.0 | 34 | 12 | III | - | - |
Prx F4 | 6.1 | 45 | ZmPrx106 | Peroxibase | 8.4 | 34 | 40 | III | - | - |