Characterization of a Lignified Secondary Phloem Fibre‐deficient Mutant of Jute (Corchorus capsularis)

Abstract

• Background and Aims High lignin content of lignocellulose jute fibre does not favour its utilization in making finer fabrics and other value‐added products. To aid the development of low‐lignin jute fibre, this study aimed to identify a phloem fibre mutant with reduced lignin.

• Methods An x‐ray‐induced mutant line (CMU) of jute (Corchorus capsularis) was morphologically evaluated and the accession (CMU 013) with the most undulated phenotype was compared with its normal parent (JRC 212) for its growth, secondary fibre development and lignification of the fibre cell wall.

• Key Results The normal and mutant plants showed similar leaf photosynthetic rates. The mutant grew more slowly, had shorter internodes and yielded much less fibre after retting. The fibre of the mutant contained 50 % less lignin but comparatively more cellulose than that of the normal type. Differentiation of primary and secondary vascular tissues throughout the CMU 013 stem was regular but it did not have secondary phloem fibre bundles as in JRC 212. Instead, a few thin‐walled, less lignified fibre cells formed uni‐ or biseriate radial rows within the phloem wedges of the middle stem. The lower and earliest developed part of the mutant stem had no lignified fibre cells. This developmental deficiency in lignification of fibre cells was correlated to a similar deficiency in phenylalanine ammonia lyase activity, but not peroxidase activity, in the bark tissue along the stem axis. In spite of severe reduction in lignin synthesis in the phloem cells this mutant functioned normally and bred true.

• Conclusions In view of the observations made, the mutant is designated as deficient lignified phloem fibre (dlpf). This mutant may be utilized to engineer low‐lignin jute fibre strains and may also serve as a model to study the positional information that coordinates secondary wall thickening of fibre cells.

INTRODUCTION

Phloem (or bast) fibres are a common source of commercial fibres and are produced from several plant species, including Corchorus capsularis and C. olitorius (jute), Linus usitatissimum (flax), Boehmeria nivea (ramie) and others (Kirby, 1963). In these dicotyledonous plants, the commercial fibres are sclerenchyma cells with copious secondary wall thickening (Kundu, 1944; Fahn, 1990). The lignin content of the fibre cell wall varies from species to species. While it is around 15 % in jute, it is less than 5 % in flax and ramie (Urquhart and Howitt, 1953; Gorshkova et al., 2000). Flax and ramie are in greater demand for producing finer fabrics. Any attempt to reduce the amount of lignin in the phloem fibre of jute, which is generally used for coarser fabrics, would be valued for potential diversification of jute usage (Palit, 1999, 2001). Unfortunately there has been no systematic effort in this direction.

Lignin, a complex phenylpropanoid polymer, is deposited in cell walls of supporting and conducting tissues, such as fibres and tracheary elements of higher plants (Wallace and Fry, 1994). It imparts rigidity to the cell wall (Lewis et al., 1999) and is responsible for decay resistance that enables the cells to withstand the saprophytic activity of organisms capable of degrading non‐lignified cell walls (Nicholson and Hammerschmidt, 1992).

From a human perspective the presence of lignin in plants has a negative impact on biomass utilization. It constrains our use of lignocellulosic materials (e.g. wood and bast fibres) in agriculture and industry. Lignins have to be removed from the wood by environmentally hazardous and costly pulping processes to manufacture paper (Ingber et al., 1985).

Potential economic benefits of modifying lignin through biotechnology have inspired many efforts to engineer lignin quantity and quality (Campbell and Sederoff, 1996). Most of the mutants and transgenics with modified lignin that have been targeted for genetic engineering are xylem mutants (Barriere and Argillier, 1993; Ralph et al., 1997; Zhong et al., 2000), at least in part because wood is more important for industrial processing. Also, the tracheary elements are easier to detect than the phloem elements (Aloni, 1988). Phenotypic screening for phloem mutants is also difficult, as lignin‐deficient sclerenchyma cells of phloem tissue do not collapse (Palit et al., 2001) as easily as in tracheary cells (Smart and Amrhein, 1985).

There are conflicting reports on plants displaying normal development even though they have large lignin deficiencies (Barriere and Argillier, 1993; Piquemal et al., 1998; Zhong et al., 1998, 2000). Additional studies are necessary, especially on woody plant species, to address these apparently contradictory effects. It is also important to investigate lignin modification in the tissue that has specific commercial demand.

This paper reports on an induced mutant of Corchorus capsularis, which has a defect in the development of the highly lignified secondary phloem fibre normally found in jute.

MATERIALS AND METHODS

Plant material

A Corchorus capsularis L. variety (JRC 212) and a true‐breeding, x‐ray (50 Kr) induced mutant (Accession number CMU 013, CRIJAF) derived from it in a radiation breeding programme in the 1950s (Anon., 1999) were grown under a polythene shed, where the environment was maintained at an ambient level. The plants were raised in glazed tile pots containing coarse, acid‐washed, neutral pH sand and were periodically supplied with complete nutrient solution (pH 7·8) and de‐ionized water. Adequate measures were taken to protect the plants from biotic and abiotic stresses that may influence lignin biosynthesis (Whetten and Sederoff, 1995). Seeds were sown on 14 Apr. and the matured plants were harvested on 16 Aug. before the onset of flowering (Ghosh, 1983).

In a preliminary study, all other accessions (CMU 010–CMU 014) stored as the same mutant line in the CRIJAF gene bank were grown in a similar manner and characterized by their morphological appearance.

Photosynthesis, growth and yield measurements

Net photosynthesis of recently matured upper leaves was measured under saturating PAR (>1000 µmol m^–2 s^–1) using a portable gas exchange system (CID, USA). Each value presented is a mean of 30 readings from six separate leaves of similar age from three separate plants of each genotype. Plant growth was determined periodically by measuring the above‐ground length of the stems of six selected plants of each genotype. Wood (stick) and fibre yields of 25 randomly harvested plants of each genotype were determined after retting in a tank (Ghosh, 1983). Daily inspection was necessary to check the end‐point of retting.

Histological procedures

To compare the vascular development at different growth stages, histological observations were made along the length of the stem. The upper and middle parts of the stem from 80‐d‐old plants were fixed in alcohol/acetic acid (3 : 1) and embedded in paraffin wax. Transverse sections were cut (10–50 µm) using a rotary microtome (Leica, Germany). The thicker, lower parts were cut (100 µm) directly using a sledge microtome (Leitz, Germany). The sections were stained with safranin and fast green according to standard protocol (Johansen, 1940). Observations were made using a Zeiss (Germany) transmitted light microscope equipped with a Nikon FM 10 camera. Kodak Gold 100 ASA film was used for photography.

Determination of lignin and lignification

Microscopy.

Transverse sections were hand‐cut from the middle part of mature (120 d old) stems and stained without fixation with phloroglucinol/HCl reagent, which specifically stains native lignins of the plant cell wall (Srivastava, 1966; Chabannes et al., 2001), and photographed under the bright field microscope.

Chemical analysis.

The chemical constituents of the matured jute fibre were determined essentially following the method of Sengupta et al. (1958). Finely milled dry jute fibres were thoroughly cleaned of wax and oils in a soxhlet apparatus using a mixture of benzene and ethanol (2 : 1 v/v) and dried under vacuum. Five grams of the sample was treated with 0·7 % sodium chlorite solution at pH 4·0, maintained by acetic acid/acetate buffer (0·2 m) for 2 h in a boiling water bath. The process was repeated three times and the samples were brought to neutral pH by washing with 2 % sodium metabisulphite and water at 60 °C, cooled and dried in P₂O₅ desiccators to a constant weight to get the holocellulose. The lignin content was determined by subtracting the amount of holocellulose from that of the de‐waxed sample.

A measured sample of holocellulose was treated with 17·5 % caustic soda at 20 °C (liquor ratio, 15 : 1) for 2–3 h. The insoluble residue was filtered through pre‐weighed gooch crucibles and washed with dilute NaOH (2 % followed by 0·2 %) and water. The filtrate was made acidic with 0·2 % acetic acid and then an equal volume of absolute ethanol was slowly added to it with constant stirring. The procedure was repeated three times and the filtrate was dried in an oven at 100 °C and cooled over phosphorus pentoxide. The amount of α‐cellulose was determined gravimetrically. The hemicellulose content was determined by subtracting the amount of α‐cellulose from that of the holocellulose sample.

For each genotype the whole fibre extracts from at least 20 individual plants were pooled and the results are the measurements of three independent analyses on random samples from the pooled material.

Measurement of fibre tenacity (tensile strength)

The tensile strength of the fibre was measured using a bundle strength tester (Bandopadhyay and Mukhopadhyay, 1964), which determined the breaking load (kg) of clean and dry fibre strands (12·5 cm length, weighing 200–300 mg) collected from near the middle portion of the fibre reeds. The tensile strength (g tex^–1) of the samples was calculated from the ratio of breaking load (kg) to its linear density (tex or g km^–1). The results are the mean and standard error of 25 fibre samples from individual plants for each genotype.

Extraction and assay of enzymes

The activity of phenylalanine ammonia lyase (PAL) (EC 4.3.1.5) and peroxidase (EC 1.11.1.7) were determined in the fibre‐rich jute bark. Bark strips peeled from upper, middle and lower parts of the stem were separately homogenized successively in phosphate buffer (pH 7·0), methanol and water to clear the chlorophyll and other intracellular contents. The thick homogenate was poured onto double layers of surgical gauge and carefully squeezed to remove the mucilage. The procedure was repeated several times in order to remove the large amount of mucilage present in jute bark. The particulate residue was re‐suspended in the same buffer and centrifuged at 2500 g for 10 min. The precipitate was taken as the cell wall fragments of phloem tissue.

The cell wall fragments were extracted either with 0·2 m borate buffer (pH 8·7) containing 5·0 mm β‐marcaptoethanol for PAL, or with 0·5 m phosphate buffer (pH 7·4) for peroxidase, using a mortar and pestle and a few grains of neutral sea sand (Sigma). The extracts were centrifuged at 5000 g for 15 min. The supernatants were saved for enzyme assays.

The PAL activity was assayed spectrophotometrically by the formation of trans‐cinnamic acid following the method of Brueske (1980) and was expressed as µmole trans‐cinnamic acid formed per µg protein. Similarly, the peroxidase activity was assayed from the oxidation of catechol by H₂O₂ in presence of the extract (Chance and Machly, 1955). A portion of the extract was used to estimate for soluble protein (Bradford, 1976) using bovine serum albumin as standard. For each genotype, five individual plants were pooled and the results represent the mean and standard error of four independent extractions and corresponding enzyme assays.

Statistical analysis of data

A t‐test was applied to determine the significance of differences in relevant characters between the two genotypes. Analysis of variance was applied only to the plant growth data (Fig. 2) to test the significant difference between genotypes and their interaction with time (Zar, 1999)

Fig. 2. Increase in plant height of JRC 212 and CMU 013 between 45 and 75 d from sowing. LSD P < 0·01 for genotype = 2·47, genotype × days = 1·75.

RESULTS

Morphology, growth and yield

Though all the accessions of the same mutant line showed morphological differences from that of their normal parent (JRC 212), CMU 013 showed maximum undulation of the stem axis, while that of CMU 014 was almost straight (Fig. 1A). The petioles and lamina of CMU 013 appear equally wavy and the stem showed shorter internodes as compared with the normal type (Fig. 1B).

Fig. 1. Phenotype of the normal (JRC 212) and mutants (CMU). A, The stem segments (m, middle; b, bottom) of (i) CMU 010, (ii) CMU 011, (iii) CMU 012, (iv) CMU 013 and (v) CMU 014. B, Portions of (i) JRC 212 and (ii) CMU 013 plants. CMU 013 shows undulation in the main axis, petiole and lamina.

In spite of the undulated phenotype, the mutant remained upright but grew significantly slower than the parent plants (Fig. 2). The differences in leaf photosynthetic rate, stomatal resistance and the rate of transpiration between the mutant and the normal type were insignificant under optimum ambient conditions (Table 1).

Table 1.

Net photosynthesis (P_N), transpiration (E) and stomatal resistance (r_s) of the leaves (± s.e.) of JRC 212 and CMU 013 grown at an average air temperature of 39·7 °C, 69 % relative humidity and 1014 µmol m^–2s^–1 PAR

Genotype	PN (µmol m–2 s–1)	E (mmol m–2 s–1)	rs (m2 s mol–1)
JRC 212	8·977 ± 1·770	3·633 ± 0·360	3·166 ± 0·618
CMU 013	9·188 ± 1·185	2·788 ± 0·390	4·388 ± 0·853
	ns	ns	ns

ns, Not significant P < 0·05.

At harvest (125 d after sowing) the mutant was significantly (approx. 31 %) shorter than the normal type (Table 2). Nevertheless, the difference in total wood weight between the normal and the mutant type was insignificant. Since the mutant plants were shorter, this insignificant difference in the amount of wood per plant indicates the production of thicker xylem elements in the mutant. The picture was reversed for phloem (bast) fibre, which was significantly less in the mutant than in the normal type. While the normal type showed 3·17 % fibre, the mutant had only 0·37 % fibre per unit of their respective stem height. There was a clear deficiency of bast fibre along the length of the mutant stem.

Table 2.

Plant height, wood weight and fibre weight per plant (± s.e.) of JRC 212 and CMU 013 at 125 d from sowing

Genotype	Plant height (cm)	Wood weight (g)	Fibre weight (g)
JRC 212	312·00 ± 7·69	20·90 ± 2·85	9·90 ± 1·01
CMU 013	216·00 ± 8·53	17·90 ± 2·07	0·79 ± 0·07
	**	ns	**

**, Significant P < 0·01; ns, not significant.

The retting or the separation of fibre strands from the rest of the stem tissue took only 7–8 d for the mutant as compared with 16–18 d for the normal stems under identical conditions.

Vascular development in the intact plant

The younger, upper part of the stems of both normal and mutant (Fig. 3A and B) did not show any secondary meristem. Thus the vascular tissues appeared to be primary in origin. The gross anatomical features, including the presence of prominent groups of primary phloem fibre bundles below the cortical zone, were also similar in both genotypes. In the comparatively mature middle region (Fig. 3C and D) a distinct vascular cambium producing conical secondary phloem wedges outside the xylem was visible in both genotypes. A clear distinction between mutant and normal plants was evident in the development of thick‐walled fibre cells within the phloem wedges. In the normal type (Fig. 3C), groups of thick‐walled cells forming successive layers of fibre bundles were separated from each other by thin‐walled parenchyma cells. The fibre bundles occupied most of the phloem tissue. In the mutant, these fibre bundles were missing (Fig. 3D). The phloem wedges were predominantly composed of thin‐walled cells, many of which were identical in their outline with the fibre cells of normal type but without the characteristic wall thickening. A few uni‐ or biseriate radial rows of thick‐walled cells were visible, mostly on the flanks of the wedges. Even these cells were absent in the lower (and consequently most mature) part of the mutant stem, which otherwise showed a larger phloem area compared with its middle part (Fig. 3F). In contrast, the uniformity of fibre bundle development was maintained in the lower stem of the normal type, which showed more layers of fibre bundles in the larger phloem wedges (Fig. 3E).

Fig. 3. Transverse sections of the stem segments of JRC 212 (A, C and E) and CMU 013 (B, D and F). A and B, Upper portions; C and D, middle portions; E and F, lower portions. pp, Bundles of primary phloem fibre cells; sp, wedge of secondary phloem; xy, secondary xylem; c, cambium. Note the deeply stained fibre bundles (fb) in the wedge of JRC 212. Bar = 100 µm.

The lower portion of the retted normal plant yielded a dense mantle of fibre (Fig. 4B), while the same portion of the mutant gave only non‐fibrous cork‐like tissue in continuation with the fibres obtained from the upper part of the stem (Fig. 4A and B).

Fig. 4. Nature of CMU 013 and JRC 212 plants after retting. A, A bunch of CMU 013 plants showing the upper fibrous and lower non‐fibrous portions. B, Magnified lower portions. Note (i) the corky nature of CMU 013 and (ii) the fibrous nature of JRC 212.

Morphology and lignification of fibre cells

The morphological comparison was made after staining the transverse sections from the middle part of the mature stems with phloroglucinol–HCl reagent. In the normal plant, while the walls of many other cells were colourless, the cells forming tight groups showed the characteristic pinkish stain for lignin (Fig. 5A). These cells in phloem wedges were secondary phloem fibre cells as they formed the typical patches of bundles in the normal type (Fig. 5A). Each fibre cell of the normal type (Fig. 5A) appeared narrower with comparatively more lignified wall than those of the mutant type (Fig. 5B–D), which showed wider cells with larger lumens and a thinner wall forming uni‐ or biseriate layers (Fig. 5B and C). Some lignified cells were also found scattered singly within the phloem wedges (Fig. 5D).

Fig. 5. Phloroglucinol stained cells in the secondary phloem wedges at the middle portion of JRC 212 (A) and CMU 013 (B–D). Bar = 10 µm.

Chemical and physical properties of the extracted fibre strands

Chemical analysis in three separate determinations showed the lignin content of the mutant fibre was almost 50 % less than the normal type (Table 3). The average lignin content in the mutant was 8·49 % while that in the normal plant was 18·34 %. In contrast, the amount of α‐cellulose, but not necessarily the hemicellulose, was nearly 30 % more in the mutant fibre than in the normal type.

Table 3.

Chemical composition (% of de‐waxed fibre) of fibre obtained from JRC 212 and CMU 013 plants grown and retted under identical conditions

Genotype	No. of determinations	α‐Cellulose	Hemicelluloses	Lignin
JRC 212	1	44·57	33·30	21·36
	2	59·33	23·08	16·59
	3	48·76	34·01	17·07
Mean		50·89	30·13	18·34
s.e.		4·40	3·53	1·52
CMU 013	1	70·91	20·96	8·12
	2	67·85	23·20	9·01
	3	60·66	29·99	8·35
Mean		66·47	24·72	8·49
s.e.		3·04	2·72	0·27

The actual values with mean and s.e. (standard error) from three independent determinations are given to show the variation among the determinations.

The tensile strength measurements of the mutant fibre showed a relatively high standard error, possibly due to the difficulty in getting uniform samples from the small amounts of extractable fibre from the mutant plants. The mean values did not differ significantly from that of the normal type (Table 4).

Table 4.

Mean tensile strength of fibre bundles (g tex^–1) ± s.e. of JRC 212 and CMU 013

Genotype	Tensile strength
JRC 212	18·94 ± 1·35
CMU 013	17·63 ± 4·85
	ns

ns, Not significant P < 0·05.

PAL and peroxidase activity in the bark (phloem tissue)

The data presented in Table 5 indicate that the activity of PAL was more or less uniform in all the stem segments of the normal type. In the mutant it was three to four times less than the normal type in the upper and middle stem, and not detectable in its lower segment.

Table 5.

Activity of phenylalanine ammonia lyase (µmol trans‐cinnamic acid formed µg^–1 protein) in the crude extract of phloem (bark) tissue of JRC 212 and CMU 013 (± s.e.)

Plant part	JRC 212	CMU 013
Upper stem	5·058 ± 0·076	1·489 ± 0·014
Middle stem	7·248 ± 0·085	1·750 ± 0·028
Lower stem	6·547 ± 0·051	Not detectable

The activity of cell wall‐bound peroxidase was more or less similar in all the stem segments of both the normal and mutant plants (Table 6).

DISCUSSION

This study shows that mutation can specifically affect the development of secondary phloem fibre in woody dicotyledonous plants without disrupting the development of other vascular tissues. It also provides genetic evidence that, in the phloem, lignin synthesis is regulated by age‐specific signals.

The defect in the mutant was most convincingly demonstrated from the finding of an adequate amount of wood (secondary xylem tissue) but a much reduced amount of phloem fibre strands upon retting of the green plants. Retting of jute involves microbial actions that disintegrate the thin‐walled cells of the bark, freeing the thick‐walled secondary phloem tissues. The jute fibre strands are manually extracted and separated from the stick or wood as soon as the process is completed (Ghosh, 1983). Jute fibres are lignocellulosic (Kirby, 1963) and thus the secondary wall development of the sclerenchymatic fibre cells involves deposition of lignin over the cellulose matrix (Preston, 1974; Grabber et al., 1991). Presence of lignin in the fibre cell wall makes it impermeable and resistant to microbial degradation (Lewis and Davin, 1994). Under similar conditions of natural microbial action the mutant yielded only a meager amount of phloem fibres as compared with that of the normal type, indicating the deficiency of lignified phloem fibre cells in the mutant.

Histological evidence also indicates that the secondary phloem fibre cells in the mutant are not only rare but that they are also not in compact groups typical of jute (Kundu, 1944). Since the cambial activity in the mutant was, in general, normal, it is difficult to believe that mutation could render specifically only those initials that differentiate the phloem fibre cells ineffective. Instead the maturation of fibre cells by thickening of their secondary walls with deposition of lignin on the cellulose matrix at the last stage (Fukuda, 1996) is suppressed, as lignification of cells is far more flexible (Sederoff et al., 1999; Ye et al., 2002). Plants with both lignified and non‐lignified fibre cells are common (Fahn, 1990). The heavy lignin content of the walls results in an increased surface area of the fibre cells, increasing cell‐to‐cell contact and leading to the Formation of groups of attached cells (Priestley and Scott, 1936; Kundu et al., 1959; Preston, 1974). Thus the lack of sufficient lignified fibre cells in the mutant might be the reason that the fibre bundles do not develop. Many industrial processes require the removal of lignin to separate the individual fibre cells (Harlley, 1987).

Whenever lignification can occur in few cells, the cells may at least be able to stick together and withstand microbial degradation (Amthor, 2003) and yield fibre strands upon retting. Yet these fibre stands showed nearly 50 % less lignin than those obtained from the normal plant. The method of lignin estimation that was followed here is mainly to isolate the total holocellulose moiety from jute fibre (Sengupta et al., 1958) and is different from that of the more widely used Klason method which employs H₂SO₄ hydrolysis. The Klason method fails to remove all non‐phenolic compounds including some carbohydrates, perhaps due to the high amount of crystalline cellulose in bast fibres, giving an overestimation of lignin (Sarkar, 1931; Gorshkova et al., 2000). The process of lignification in the mutant fibre wall must have uncoupled to a great extent from the development of the cellulose matrix. Cellulose was more than adequate in giving the mutant fibre a good tensile strength. This is possibly due to the fact that dehydrated cellulose is a far stiffer cell wall constituent (Niklas, 1989) and the strength of the dry jute fibre is little affected after removal of lignin (Sarkar, 1935).

The basal portion of the mutant was totally devoid of lignified fibre cells, while at least a few phloem cells in the middle and upper part of the stem were lignified and yielded some amount of hard fibre. This indicates that the signal for lignification of secondary cell walls is not uniformly operative at different developmental stages of the vascular plant. The mutant provides genetic evidence that the signal may remain blocked in the secondary phloem cells developed early in the plant’s life but starts to function in the middle and upper regions, which developed later.

This observation of age‐induced lignification was consistent with the expression of PAL activity, which is one of the key regulators for biosynthesis of phenylpropanoid compounds including lignin (Northcote, 1995; Anterola and Lewis, 2002). PAL activity was recorded only in the middle and upper part of the mutant stem. The comparatively lower PAL activity in the mutant than in the normal plant suggests a block upstream in the shikimate pathway, lowering its lignin synthesis in the phloem tissue. On the other hand, the activity of peroxidase, one of the downstream enzymes assigned to the oxidative polymerization of monolignols in many plants (Lagrimini, 1991; Nose et al., 1995), was more or less similar in the mutant and the normal type. This suggests that this oxidative enzyme might have a non‐specific role in lignin biosynthesis in jute, as in the cases reported for other plants (Lewis and Yamamoto, 1990; Chabbert et al., 1992).

Despite normal photosynthesis, the reduced growth rate and abnormal phenotype of the mutant also points to a defective metabolism of phenolic compounds, which have been assumed to interfere with the growth of the plant by altering the function of plant growth substances (Chabannes et al., 2001). Some of these, especially IAA and GA₃, have profound effects on differentiation and growth of phloem fibres (Aloni, 1988; Aloni et al., 1990). The observed down‐regulation of PAL in the mutant might have induced a pleiotropic effect on its growth. Elkind et al. (1990) found similar diverse effects like an unusual phenotype, an altered leaf shape, stunted growth and other features by down‐regulation of phenylpropanoid biosynthesis in transgenic tobacco using a heterologous phenylalanine ammonia lyase gene.

In view of these findings, CMU 013 is considered to be a pure line jute mutant with undulated phenotype and distinct deficiency in lignified secondary phloem fibre, all due to down‐regulated lignin biosynthesis in the secondary phloem cells. The mutant is designated as deficient lignified phloem fibre (dlpf) and it is anticipated that it will be utilized to genetically engineer low‐lignin jute fibre, for which there has been a long‐standing demand in jute‐based industries (Chatterjee et al., 1991).

ACKNOWLEDGEMENTS

We thank Dr H. S. Sen, Director, CRIJAF for his keen interest in this work and Dr B. C. Sasmal for helping in the statistical analysis of data. The Indian Council of Agricultural Research (ICAR), AP Cess Fund grant 9‐12/96‐CCI, supported this work.

Table 6.

Activity of cell wall‐bound peroxidase (OD change min^–1 mg^–1 protein) in the crude extract of phloem (bark) tissues of JRC 212 and CMU 013 (± s.e.)

Plant part	JRC 212	CMU 013
Upper stem	0·403 ± 0·01	0·419 ± 0·012
Middle stem	0·490 ± 0·011	0·402 ± 0·016
Lower stem	0·479 ± 0·015	0·419 ± 0·011

Received: 23 June 2003; ; Returned for revision: 22 September 2003; Accepted: 27 October 2003    Published electronically: 5 January 2004