DISTRIBUTION O F CHROM IUM AND ITS EFFECT ON M ORPHOLO GY AND
ANATOM Y OF BRASSICA JUNCEA (INDIAN MUSTARD) AND SOIL
M ICROORGANISM S
by
Svetlana Bluskov
B.Sc., Rostov-on-Don State University, 1994

THESIS SUBM ITTED IN PARTIAL FULFILLMENT OF
THE REQUIREM ENTS FOR THE DEGREE OF
M ASTER OF SCIENCE
in
NATURAL RESOURCES AND ENVIRONM ENTAL STUDIES
(ENVIRONM ENTAL SCIENCE)

THE UNIVERSITY OF NORTHERN BRITISH COLUMBIA
August, 2004

© Svetlana Bluskov, 2004

1^1

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ABSTRACT

The present study was conducted to evaluate the growth response of
Brassica juncea (Indian mustard) and soil microorganisms to two forms of
chromium

(Or)

and to study mechanism(s)

involved

in Or binding

and

sequestration by the plant. The chemical spéciation of Or in rhizosphere soil was
also investigated in this thesis.

S. juncea was grown under greenhouse conditions in field-moist or airdried soils, amended with either Or (III) or Or (VI). The plant concentrated
approximately 200 mg Or per kg of root dry weight (DW) and 6 mg Or per kg of
shoot DW in the Or (lll)-amended soil.

In the Or (Vl)-amended soil, the plant

accumulated approximately 390 mg Or per kg of root DW and 20 mg Or per kg
of shoot DW. In general, the plant was tolerant of both Or treatments. Or (VI)
appeared to be more toxic to both soil microorganisms and plant morphology
and anatomy than Or (III) and root growth and anatomical characteristics were
inhibited to a greater extent than those of shoots. Soil chemical analyses and Xray absorption near-edge spectroscopy (XANES) detected Or (III) species in Or
(VI) treatment. The XANES and X-ray microprobe spectroscopy data for the
plant tissues revealed nearly complete conversion of Cr (VI) to Cr (III) in the
roots, where it was accumulated preferentially in epidermal and cortical cells as
Cr (lll)-acetate (72%), while, in the leaves, it was concentrated in epidermal and
spongy mesophyll cells as Cr (lll)-oxalate (81%). The ability of B. juncea to
tolerate and detoxify Cr (VI) within the roots makes this plant a potential
candidate for phytostabilization.

11

TABLE OF CONTENTS

A B S T R A C T............................................................................................................................ii
TABLE OF C O N T E N T S .....................................................................................................iii
LIST OF ABBREVIATIONS AND S Y M B O L S ............................................................... vi
LIST OF F IG U R E S ............................................................................................................vill
LIST OF T A B LE S ................................................................................................................. x
ACK N O W LEDG EM ENTS................................................................................................. xii
1.0.

IN T R O D U C T IO N .......................................................................................................1

2.0.

BACKGROUND........................................................................................................ 7

2.1.

Distribution of Chromium in Soil........................................................................ 7

2.1.1. Sources of soil chromium..............................................................................7
2.1.2. Chromium spéciation, mobility and availability......................................... 8
2.2.

Chromium and Soil Microorganisms............................................................. 12

2.2.1. Chromium inhibitory effects........................................................................ 12
2.2.2. Microbial reduction of Cr (V I)...................................................................... 13
2.3.

Heavy Metals and Plants................................................................................. 14

2.3.1. Metal toxicity................................................................................................... 14
2.3.2. Mechanisms of metal uptake...................................................................... 17
2.4.

Brassica ju n cea.................................................................................................. 19

2.4.1. Geographical distribution............................................................................. 19
2.4.2. U s e ................................................................................................................... 19
2.4.3. Life cycle..........................................................................................................20
2.4.4. Growth requirements....................................................................................21

111

2.4.4.1. Soil type....................................................................................................21
2.4.4.2. Soli fertility............................................................................................... 21
2.4.4.3. Tem perature............................................................................................22
2.4.4.4. W ater.........................................................................................................22
2.4.5. Heavy metal uptake...................................................................................... 23
2.4.6. Heavy metal phytotoxicity........................................................................... 24
2.4.6.1. Effects on shoots....................................................................................24
2.4.6.2. Effects on roots....................................................................................... 25
2.5.

Chromium and Plants........................................................................................ 26

2.5.1. Chromium impact on plant growth and development............................ 26
2.5.1.1. Beneficial effects.....................................................................................26
2.5.1.2. Visible deleterious effects

......................................................... 27

2.5.1.3. Cr (III) and Cr (VI) phytotoxicities........................................................28
2.5.1.4. Morphological and anatomical alterations.........................................29
2.5.2. Chromium absorption and distribution......................................................31
3.0.

MATERIALS AND M E TH O D S .............................................................................34

3.1.

Soil Source and Preparation............................................................................ 34

3.2.

Chromium Application....................................................................................... 35

3.3.

Plant M aterial.......................................................................................................37

3.4.

Plant Growth and HarvestConditions............................................................. 37

3.5.

Soil Analyses........................................................................................................38

3.5.1. Soluble Cr (III) and Cr (V I).......................................................................... 38
3.5.2. Microbial biomass C .....................................................................................39
3.5.3. Chromium spéciation in soils......................................................................41
3.6.

Plant A nalyses.................................................................................................... 42

3.6.1. Plant growth and chemical measurements.............................................42
3.6.1.1. Visual evaluation of Cr stress..............................................................42
3.6.1.2. Plant height.............................................................................................. 42

IV

3. 6 .1.3. Shoot and root dry weight.....................................................................43
3 .6 .1.4. Chromium accumulation.......................................................................43
3.6.1.5. Low-molecular-weight organic acids in root exudates....................44
3.6.2.

Anatomical measurements....................................................................... 45

3.6.2.1. Stem anatomical characteristics......................................................... 45
3.6.2.2. Root and leaf anatomical characteristics.......................................... 46
3.6.2.3. Chromium spéciation in plant tissues.................................................47
3.6.2 4. Chromium localization within plant..................................................... 48
3.7.

Statistical A nalyses.......................................................................................... 49
RESULTS AND D IS C U S S IO N ............................................................................50

4.0.
4.1.

Soluble Cr (III) and Cr (VI) in so il.................................................................. 50

4.2.

Chromium spéciation in soil............................................................................54

4.3.

Chromium influence on soil microbial activity............................................. 60

4.4.

Macroscopic effects of chromium on plant grow th.................................... 64

4.4.1.

Chromium uptake........................................................................................ 64

4.4.2.

Chromium influence on plant visible stress............................................70

4.4.3.

Chromium influence on plant root and shoot growth...........................72

4.4.4.

Chromium influence on plant root exudation.........................................77

4.5.

Microscopic effects of chromium on plant growth.......................................8 8

4.5.1.

Chromium influence on shoot anatomical characteristics.................. 8 8

4.5.2.

Chromium influence on root anatomical characteristics..................... 92

4.5.3.

Chromium spéciation and distribution within plant................................99

5.0.

C O N C L U S IO N S ................................................................................................... 109

6.0.

FUTURE STUDIES AND R EC O M M EN D A TIO N S........................................ 112

7.0.

R E F E R E N C E S ..................................................................................................... 113

A P P E N D IX ........................................................................................................................ 132

LIST OF ABBREVIATIONS AND SYMBOLS

%

percent

°

degrees

°c

degrees Celclus

M9

microgram

ml

microliter

|jm

micrometer

|j M

micromolar

Alp and Pep

extractable with sodium pyrophosphate

CEO

cation exchange capacity

cm

centimeter

cmolc

centimoles of charge

Eh

redox potential

g

gram

h

hour

ha

hectare

ID

internal diameter

keV

kilo electron volts

kg

kilogram

kV

kilovolt

L

liter

M

molar

Me

metal

meq

milliequivalent

mg

milligram

min

minutes

mL

milliliter

mm

millimeter

VI

mM

millimolar

mm^

millimeter square

n

sample size

nm

nanometer

P

probability

psi

pounds per square inch

rpm

revolutions per minute

sec

second

spp.

species

a

significance level

vu

LIST OF FIGURES

Figure 3.1. Rhizotron for growing Brassica juncea....................................................36
Figure 3.2. Longitudinally-sectioned lateral root (200x) from control Brassica
Juncea showing box used for measurements of cells in a partof xylem .....48
Figure 4.1. Water-soluble Cr (III) and Cr (VI) extracted from the Cr (III) and Cr
(Vl)-amended (100 mg kg'^ of either CrCb BHaO or K2 Cr2 0 7 ) field-moist and
air-dried rhizosphere and bulk soils of Brassica Juncea after 17, 36, and 69
days of growth...........................................................................................................51
Figure 4.2. X-ray absorption near-edge spectroscopy (XANES) spectraof Cr (III,
VI) reference compounds........................................................................................55
Figure 4.3. X-ray absorption near-edge spectroscopy (XANES) spectra of fieldmoist: (a, b) Cr (lll)-amended rhizosphere and bulk soils; (c, d) Cr (VI)amended rhizosphere and bulk soils.................................................................... 57
Figure 4.4. X-ray absorption near-edge spectroscopy (XANES) spectra of airdried: (a, b) Cr (lll)-amended rhizosphere and bulk soils; (c, d) Cr (VI)amended rhizosphere and bulk soils....................................................................58
Figure 4.5. Effect of two chromium species [Cr (III) and Cr (VI)] on shoot height
of Brassica Juncea grown for 69 days.................................................................. 73
Figure 4.6. Effect of two chromium species [Cr (III) and Cr (VI)] on mean root
and shoot dry weight of Brassica Juncea grown for different exposure
periods.........................................................................................................................75
Figure 4.7. Electropherograms of: (a) 10 pM standard solution mixture of malic,
citric, succinic and acetic acids; (b) root exudates collected from Brassica
Juncea at 17 days (electrokinetic injection of 10 kV for 10 sec)..................... 78
Figure 4.8. Electropherograms of root exudates collected from Brassica Juncea
at: (a) 36 days; (b) 69 days (electrokinetic injection of 10 kV fo r 10 s e c )... 79
Figure 4.9. Electropherograms of: (a) 100 pM standard solution mixture of malic,
citric, succinic and acetic acids; (b) root exudates collected from Brassica
Juncea at 17 days (pressure injection of 0.1 psi for 10 sec)............................ 82

V lll

Figure 4.10. Electropherograms of root exudates collected from Brassica juncea
at: (a) 36 days; (b) 69 days (pressure injection of 0.1 psi for 10 sec)........... 83
Figure 4.11. Effect of chromium on tap root anatomy...............................................95
Figure 4.12. Effect of chromium on lateral root anatomy......................................... 98
Figure 4.13. XANES spectra and LC-XANES fittings of Cr (Vl)-treated leaf and
root of Brassica juncea grown for 69 days in the field-moist soil................. 100
Figure 4.14. X-ray microprobe image (200x) of a cross-sectioned leaf of
Brassica juncea grown for 69 days in field-moist soil treated with 100 mg
kg-1 CrCl3 6 H 2 0 ........................................................................................................105
Figure 4.15. X-ray microprobe image (200x) of a longitudinal-sectioned lateral
root of Brassica juncea grown for 69 days in field-moist soil treated with 100
mg kg'i CrCb OHaO................................................................................................ 106

IX

LIST OF TABLES

Table 3.1. Physical and chemical properties of soil from Aleza Lake................... 35
Table 4.1. Distribution of chromium compounds (%) in the rhizosphere and bulk
field-moist and air-dried soils of Brassica juncea treated with 100 mg kg'^of
either CrCb 6 H 2 O or K2 Cr2 0 7 ................................................................................. 59
Table 4.2. Chloroform fumigation-extraction C flush (mg 0 kg'^ D W soil) in fieldmoist and air-dried bulk and rhizosphere soils of Brassica juncea treated
with 1 0 0 mg kg'^ of either CrCl3 .6 H 2 0 or K2 Cr2 0 7 .............................................61
Table 4.3. Chromium concentration in roots and shoots of Brassica juncea after
69 days of growth in Cr (III, VI)-contaminated field-moist and air-dried soils
..................................................................................................................................... 65
Table 4.4. Chromium accumulation in roots and shoots of Brassica juncea after
69 days of growth in Cr (III, VI)-contaminated field-moist and air-dried soils.

66
Table 4.5. Visible stress in Brassica juncea after 69 days of growth in Cr (III, VI)contaminated field-moist and air-dried soils....................................................... 71
Table 4.6. Reproducibility of pressure sample introduction of 100 pmol L"^
standard mixture of malic, citric, succinic, and acetic acids in capillary
electrophoresis.......................................................................................................... 81
Table 4.7. Organic acids (pg L'^) in root exudates of Brassica juncea treated with
100 mg kg'^ of either CrCl3 6 H 2 0 or K2 Cr2 0 7 ......................................................85
Table 4.8. Effect of two chromium species [Cr (III) and Cr (VI)] on number of
vascular bundles and xylem cells, stem diameter, and width of epidermis,
cortex, phloem, xylem, and pith in Brassica juncea after 69 days of growth
..................................................................................................................................... 89
Table 4.9. £/fecf of two chromium species [Cr (III) and Cr (VI)] on leaf thickness,
thickness of palisade and spongy mesophyll, palisade cell layer number,
and leaf vein number of Brassica juncea after 69 days of growth................. 91
Table 4.10. Effect of two chromium species [Cr (III) and Cr (VI)] on root
diameter, xylem diameter, number and width of all cells in a part of xylem

X

and large cells in entire xylem of tap and lateral roots of Brassica Juncea
after 69 days of growth............................................................................................93
Table 4.11. Distribution of chromium compounds (%) in a root and leaf of
Brassica juncea grown for 69 days in field-moist soil treated with 100 mg
kg-^ of KzCrzOy.........................................................................................................103
Table 1 (Appendix). Soil profile from Aleza Lake..................................................... 132

XI

ACKNOW LEDGEM ENTS

I would like to express my gratitude to my supervisor, Dr.

Joselito M.

Arocena, for giving me the opportunity to be one of his students, do research in
his laboratory, learn, and experience throughout my graduate studies. His
continued financial support (from NSERC and CRC Program) and help with the
collection of the essential X-ray absorption spectroscopy and X-ray microprobe
data are also highly appreciated. In addition, I am grateful to the other members
of my advisory committee. Dr. Jane Young and Dr. Oladipo Omotoso. Thank you
very much for your time, extremely useful suggestions and generous assistance.

Special thanks to Dr. Hugues B. Massicote for his valuable advice on
plant roots and to Dr. Ming Chen for his preparation of root and leaf sections.
Many thanks to Dr. Syed

Khalid from X18B

beam-line at the

National

Synchrotron Light Source at Brookhaven National Laboratory and Dr. Robert A.
Gordon from 20-ID /PN C-C AT beam-line at the Advanced Photon Source at
Argonne National Laboratory for their assistance with the collection of the
XAS/XANES and X-ray microprobe data. In addition, I would like to thank the
staff of USDA/ARS Plant Introduction Station of Iowa University for their
generous supply with Brassica Juncea seeds. I must also thank the staff of the
Enhanced Forestry Laboratory, John Orlowsky and Steve Storch, for their
technical support. Dr. Dave Dick and Allen Essler for their help in collecting

XU

some analytical data, and Collin Chisholm for his computer expertise. Moreover,
I am thankful to my classmate, Geoffrey Odongo for helping me a lot with soil
collection and preparation.

My heartfelt thanks to my family, especially to my mom Lidiya and my
sister Ludmila, for their love and support, which helped me a lot throughout my
studies. Lastly, I am very grateful to my husband, Iliya, for his endless
understanding, patience, encouragement, and for always being there for me.
Thank you all.

x in

1.0.

INTRODUCTION

Soils naturally contain heavy metals as a result of proximity to mineral
outcrops or ore bodies and/or anthropogenically as a result of industrial activities
(Baker et al., 1994a). Chromium is a major soil contaminant due to its use in
metallurgy, tanneries, leathers, dyes, textile, and wood preservation (Adriano,
1986). In soils, Cr is present in two stable oxidation states: Cr (III) and Cr (VI). Cr
(III) exists in soil mostly as insoluble oxides and hydroxides. It is also associated
with soil minerals and organic matter. Therefore, it is considered stable in soils.
However, the potential for oxidation of Cr (III) to Cr (VI) can make the former
almost as hazardous as the latter.

Contrary to Cr (III), Cr (VI) is more soluble and hence mobile in soils. As a
result, Cr (VI) can enter the food chain through either leaching into groundwater
or absorption by plants.

In addition, both Cr (III) and Cr (VI) produce

carcinogenic activity in animals, including humans (Norseth, 1981). In particular,
both Cr forms can inhibit synthesis of deoxyribonucleic acid (DNA) (Tamino at
a!., 1981). Cr (VI) crosses cell membranes more easily than Cr (III) (Arslan at a!.,
1987). However, once in the system, the reduction of Cr (VI) to Cr (III) or Cr (V)
takes place (Norseth, 1981; Micera and Dessi, 1988; Liu at a/., 1995). Formed
Cr (III) and Cr (V) complexes react with hydrogen peroxide (H 2 O 2 ), generating
the highly toxic free hydroxyl ( OH) radicals, which trigger DNA oxidative damage

(Shi and Dalai, 1990). For all of the above-mentioned reasons, there is concern
relating to soil pollution issues which justifies the importance of cleaning up Cr
(III, VI)-contaminated soils.

Several methods have been used to treat metal-contaminated soils
including landfilling (excavation, transport and deposition of contaminated soil in
a permitted

hazardous waste landfill), solidification/stabilization or fixation

(chemical processing of soil to immobilize metals), and soil flushing and washing
or leaching (use of acid solutions or other leachants to desorb and leach metals
from soil) (Salt et al., 1995). However, these technologies are extremely
expensive and very disruptive to the site and the environment (Lasat, 2002).
Phytoremediation (use of plants in remediation) has been employed for years as
an alternative to the above approaches. It has become one of the most popular
techniques used in remediation because plants can be aesthetically pleasing,
economically viable, and non-destructive to the environment. Another advantage
of phytoremediation is the potential recovery of the valuable metal components
from the contaminated biomass through repeated harvests (Chaney et a!.,
1997). However, one of the drawbacks of phytoremediation is the potential
contamination of the food chain via wildlife.

The phytoremediation concept is based on the ability of particular plants
to extract, concentrate, and/or degrade contaminants (Baker and Brooks, 1989).

According to Reeves et al. (1995), hyperaccumulators are those plants which
leaves can accumulate more than 100 mg cadmium (Cd) or selenium (Se) kg'^
(DW), more than 1,000 mg lead (Pb), nickel (Ni), cobalt (Co), Cr, or copper (Cu)
kg'^ (DW), and more than 10,000 mg iron (Fe), manganese (Mn), or zinc (Zn)
kg"^ (DW) when grown in metal-rich soils. Heavy metal resistant plants can be
planted and left to grow for several seasons before they are harvested,
incinerated and the ash safely disposed of (Sellers, 1998).

To be effective remediators of soil heavy metals, plants must also have
the ability to metabolize metals and

maintain growth and

development.

Hyperaccumulators, unlike sensitive plants, have developed unique (metal and
plant-specific) resistance mechanisms to defend their cellular activity and
structures against metal stresses. These mechanisms include metal chelation
with appropriate high-affinity ligands (preferentially low-molecular-weight organic
acids originating from root exudates), biotransformation with reductants and
subsequent compartmentalization either in the cytoplasm or in a sub-cellular
organelle, i.e., the vacuole (Salt at a/., 1998). Nevertheless, further knowledge
on how these
translocation

mechanisms

function

phytoremediation.

is

and the
required

mechanisms of metal
to

enhance

the

uptake and

effectiveness

of

Brassica juncea (Indian mustard) Is a member of Kingdom Plantae
(plants),

Subkingdom

Tracheobionta

(vascular

plants),

Superdivision

Spermatophyta (seed plants). Division Magnoliophyta (flowering plants). Class
Magnoliopsida (dicotyledons). Subclass Dilleniidae, Order Capparales, and
Family Brassicaceae (mustard). It is referred to as brown mustard or Indian
mustard (USDA

NRCS,

2003). B. juncea has been widely employed in

phytoremediation because of its remarkable capacity to accumulate high levels
of heavy metals, including Ni, Zn, Cu, and Pb. For example, its shoots have
been reported to take up 308,600 mg Ni kg'^ plant (DW) and 172,300 mg Zn kg'^
plant (DW) in a fertilized sand/Perlite mixture supplemented with 100 mg Me kg'^
soil (Kumar et a!., 1995). 6 . juncea is also considered a Cr hyperaccumulator.
For example, in a study by Salt et al. (1995), when grown in a hydroponic culture
system and relatively low Cr concentration, i.e., 0.4 mg L '\ plant seedlings
accumulated 2,194 mg kg'^ DW in their roots in a single cropping. Once taken
up by the plant, Cr is generally retained in roots rather than in shoots. In a study
by Shahandeh and Hossner (2000), 6 . juncea, supplied with 100 mg Cr (III) or
Cr (VI) kg"^ soil, accumulated 191 and 4.6 mg Cr (III) kg'^ and 431 and 44.7 mg
Cr (VI) kg'^ in plant roots and shoots, respectively. Other benefits of using this
species in phytoremediation is its ability to grow quickly (matures 8 - 1 0 weeks
after sowing) and to produce a high amount of biomass in relatively short
periods of time (18 t ha'^ in 2.5 months) (Blaylock eta!., 1997).

The effectiveness of B. juncea in removal of Cr from polluted soils greatly
depends both on Cr bioavailability and the ability of the plant to tolerate the
metal. However, information concerning the Cr toxicity and metal sequestration
in this species is limited. The present study was undertaken to gain a better
general understanding of the mechanism(s) involved in Cr binding and tolerance
by plants. A further goal of this research was to examine the effects of Cr on soil
microbial activity as well as to investigate growth and developmental responses
of 6 . juncea when grown in different Cr and soil treatments. The rhizosphere
zone of the plant (soil-root interface), which serves as the main entry point of
metals into roots, was of special interest. The particular objectives of this study
were: (i) to determine the effect of Cr (III, VI) and soil (field-moist, air-dried)
treatments on solubility and, hence, availability of Cr to the plant; (ii) to assess
spéciation of Cr in polluted rhizosphere and bulk soils; (iii) to establish whether
Cr affects activity of soil microorganisms; (iv) to determine the extent to which B.
juncea can absorb Cr and to identify the form of accumulated and translocated
metal, its chemical spéciation and distribution within the plant; (v) to observe any
Cr toxicity in the plant at macroscopic (visible stress and root and shoot growth)
and microscopic (stem, root, and leaf cells) levels; and (vi) to evaluate how lowmolecular-weight organic acids in root exudates of B. juncea are influenced by
different developmental stages of the plant and Cr application. It is hoped that
knowledge gained from this study will help to elucidate the role of plants in soil

remediation and, additionally, to potentially provide knowledge of particular
heavy metal species that are detrimental to plant growth and development.

2.0.

BACKGROUND

2.1.

Distribution of Chromium in Soil

2.1.1. Sources of soil chromium

Chromium naturally occurs in minerals and rocks and can be released
into the soil environment by their weathering and erosion (Adriano, 1986).
Minerals such as chromite (FeCr 2 0 4 ) and crokoite (PbCr 0 4 ) contain high
amounts of Cr; however, they are highly insoluble and, thus, sparse in the soil
environment. Soils naturally high in Cr can be found where parent material is
derived from serpentine rocks. In particular, the concentration of Cr ranges from
5 to 1,000 mg kg'^ in natural soils, while it can reach 125,000 mg kg'^ in
serpentine soils (Adriano, 1986).

Anthropogenic inputs of Cr into soils can be a result of both industrial
activities, such as production of alloys, leather tanning, electroplating, and
dyeing, and agricultural sources, including application of pesticides, fertilizers,
and manures (Nriagu and Kabir, 1995). Moreover, improper handling and
storage of Cr-containing wood preserves can result in the transfer of large metal
quantities to local soils (Bamwoya et al., 1991). Weathering of treated lumber
exposed to acidic rainfall can also release Cr (Warner and Solomon, 1990). It is
estimated that wastes containing more than 5,000 ton of Cr in various forms are
dumped annually onto Canadian soils (The National Contaminated Sites

Remediation Program, 1996). Since Cr is not biodegradable, it has infinite
persistence in the soil environment (Bartlett, 1991).

2.1.2. Chromium spéciation, mobility and availability

Chromium

is

a

unique

heavy

metal,

because

it exists

in

soil

environments in two stable valence states, i.e., Cr (III) and Cr (VI), with
contrasting solubility, mobility, and, consequently, toxicity (Bartlett and James,
1988). The behavior of these two forms of Cr in the soil environment strongly
correlates with their potential for precipitation/dissolution, sorption/desorption,
and oxidation/reduction.

The main species of Cr (III) in soils are Cr^"" at pH < 3.6, [Cr(H 2 0 )6 ]^^ and
[Cr(H 2 0 ) 5 0 H p at pH 4,

Cr(0 H )3 at pH from 6.3 to 11.5, and [Cr(0 H)4 ]’ at pH

above 12 (Rai et al., 1989). Cr (III), like other first-row transition metals, has a
strong

tendency

to

form

octahedral

coordination

compounds,

including

complexes and chelates with oxygen (O), sulfur (S), and nitrogen (N)-containing
ligands (Nakayama eta!., 1981). As a result, Cr (III) solubility in soils is restricted
due to formation of Cr oxides,

hydroxides, and phosphates and

mixed

complexes with Fe. Moreover, Cr (III) greatly binds to clay particles and soil
organic matter and, as the pH increases, Cr (III) adsorption increases (Griffin et
a!., 1977) only for clay-sized minerals containing pH-dependent surface charges
such as the oxides of Fe, AI and Si, but not for phyllosilicates (Brady and Weil,

2002). Therefore,

Cr (III) Is considered Immobile, unavailable and, thus,

relatively harmless In soils.

Under certain soil conditions, however, Cr (III) may be oxidized to Cr (VI),
which may leach from the soil system, thereby leading to serious environmental
consequences (Bartlett and James, 1979). According to Cary (1982), oxygen
does not react appreciably with Cr (III). The most likely oxidant of Cr (III) In soil Is
Mn (IV) oxide acting as an electron acceptor and a link between Cr (III) and the
oxygen In the atmosphere (Cary, 1982; Bartlett and James, 1988). Although soil
microorganisms do not appear to be required for the oxidation (Ross and
Bartlett, 1981), they might be Involved Indirectly, because all mlcroblallymedlated transformations In soil seem to result In pH and Eh shifts (Bartlett,
1986a). The oxidation of Cr (III) by soil manganese oxides Is controlled by the
surface characteristics of the oxides and the availability of the Cr (III) to the
surface (Bartlett and James, 1988). Manganese oxides typically accumulate on
the surfaces of clays and Fe oxides (Bartlett and James, 1979). Manganese
minerals (oxides, silicates, and carbonates) have large surface areas and tend
to have negative charges at all but acidic conditions. These properties are
associated with their high absorptive capacities, particularly for heavy metals
(Bartlett and James, 1979).

Some adsorbed Cr (III) species, tightly bound to humic substances or
recently precipitated Cr hydroxides, can be readily extracted by salts of lowmolecular-welght organic acids that appear to be the most likely vehicles by
which Cr (III) moves toward oxidizing Mn surfaces (Bartlett, 1986b). The
oxidation rate of organic Cr (III) compounds Is slower than that of freshly
precipitated Cr (III) (James and Bartlett, 1983). The reason for this Is the
presence of extraneous organic materials, containing carboxyl groups and
phosphates, which can form strong complexes with Mn (III) and cause reverse
dismutatlon to take place via the following reaction;

Mn^^ + Mn02 + 4H^ ^ 2Mn^^ + 2 H2O
Mn (III) can further oxidize the organic carbon (C) that It Is holding and become
reduced to Mn (II), thereby making Cr (III) oxidation Impossible (Bartlett, 1986a).

The chemistry of Cr (VI) Is very different from that of Cr (III). Cr (VI) forms
a number of anions such as HCr 0 4 ‘ (hydrochromate), Cr2 0 7 ^‘ (dichromate) and
Cr0 4 ^' (chromate), the last being the predominant form at pH > 6 (Ral et al.,
1989). Cr (VI) compounds are very unstable and soluble In both acid and
alkaline soils (Adriano, 1986). They do not readily adsorb to surfaces. If they do,
mineral solids that contain

Inorganic hydroxyl groups.

Including

Fe and

aluminum (Al) oxides, kaollnlte, and montmorlllonlte, are among the main
adsorbents of Cr (VI) (Ral at a!., 1989). Adsorption Increases with decreasing pH
as a result of protonation of the surface hydroxyl sites (Ral at a!., 1989). The

10

intensity of adsorption to soil oxides and clay particles also depends on the
presence of competing ligands, such as phosphate, carbonate and sulfate
(Adriano, 1986). Therefore, liming and application of phosphate fertilizers, for
example, are practiced to remobilize adsorbed Cr (VI) (Bartlett and James,
1988).

The Cr (VI) ion is a stronger oxidizing agent than Cr (III) because of its
large positive reduction potential and tetrahedral coordination (Nriagu and Kabir,
1995). The high oxidizing potential and solubility make Cr (VI) more mobile and,
hence, toxic than Cr (III). On the other hand, the high oxidizing potential of Cr
(VI) means it is easily reduced to Cr (III), when electron donors, such as soil
humic compounds

(fulvic and

humic acid),

non-humic substances

(low-

molecular-weight organic acids, carbohydrates and proteins), aqueous inorganic
species (Fe (III) and sulfides) and common soil minerals (kaolinite and
montmorillonite) are present in soil (Bartlett and Kimble, 1976; Cary, 1982).
Another factor that affects the rate and extent of Cr (VI) reduction is pH.
Generally, because hydrogen is consumed in the reduction reaction, it mainly
occurs in acid soils and under both aerobic and anaerobic conditions. In
addition, soil microbial activity may influence Cr (VI) mobilization both directly via
release of special enzymes-reductases (Salt et al., 1998) or indirectly via
production of chelates, including low-molecular-weight organic acids (Bartlett,

11

1991) or depletion of soil oxygen levels which tends to lower soil pH (Losi et al.,
1994).

2.2.

Chromium and Soil Microorganisms

2.2.1. Chromium inhibitory effects

Probably because of its low solubility and high tendency to undergo
precipitation and complexation reactions, Cr (III) is not readily taken up by
microbial cells compared with the more mobile Cr (VI) that can easily penetrate
cell membranes (Arslan at a!., 1987). Therefore, the former is not considered to
be particularly toxic to soil microorganisms, while the latter is. Once inside the
cytoplasm, Cr (VI) is reduced, but its toxicity most likely results from oxidation of
cell components (Arslan at a!., 1987). However, the ability of Cr (III) to become
oxidized and produce Cr (VI), although for a short time, might be responsible for
inhibitory effects of this form of Cr on soil microbial activity (Bartlett and James,
1988). For example, Ross at al. (1981) reported that addition of 10 and 100 mg
Cr (VI) kg‘^ soil and 100 mg Cr (III) kg'^ soil significantly reduced carbon dioxide
(CO 2 ) evolution when added to loam and fine sandy loam soils. Moreover, in
spite of the fact that extractable Cr (VI) disappeared for the 3-week duration of
the study, the CO 2 evolution did not increase (Ross at al., 1981). The results of
other short- as well as long-term experiments indicate that other effects including
species abundance, nitrogen biotransformation, and enzyme (phosphatase.

12

sulphatase, arylsulphatase, and urease) activities are also inhibited in soil
microbes, when the added concentrations are 25 to 100 mg Cr (III) kg"^ soil and
1 to 10 mg Cr (VI) kg'^ soil (Tabatabai, 1977; Bollag and Barabasz, 1979;
Doelman and Haanstra, 1984, 1986; Yadav et al., 1986; Speir et a i, 1995).

2.2.2. Microbial reduction of Cr (VI)

Bacteria that are able to reduce Cr (VI) to Cr (III) have been reported by
several researchers (Horitsu et a i, 1987; Bopp and Ehrlich, 1988; Wang et a i,
1989). Representatives of the genera Aeromonas, Escherichia, Pseudomonas,
and Enterobacter are among microorganisms capable of Cr (VI) reduction
(Kvasnikov et a i, 1988). The reduction is carried out either by cell membranes
or by soluble proteins (Chen and Hao, 1998). For example, membrane-bound Cr
(VI) reductases have been found in Pseudomonas fluorescens (Bopp and
Ehrlich, 1988) and Enterobacter cloacae (Wang et a i, 1989). Such enzymatic
mechanisms ensure that this process takes place at the surface of bacterial
cells, where Cr (III) hydroxides precipitate, thereby protecting cells from the
toxicity of Cr (VI) (Wang et a i, 1989). In addition, depending on growth
conditions and type of Cr (Vl)-reducing bacteria, some organic substrates
including amino acids (peptone, glucose and lactose), salts of organic acids
(propionate, acetate, malate, succinate, citrate, pyruvate, and lactate), ethanol,
and glycerol, might serve as electron donors for the Cr (VI) reduction (Chen and
Hao, 1998). However, reduction of Cr (VI) is generally slower than its uptake in

13

bacterial systems

(Ohtake

and

Silver,

1994);

as a

result,

in order for

detoxification to occur, bacteria must be able to tolerate the toxicity of Cr (VI). It
has been demonstrated that Cr (VI) resistance is plasmid-associated. The genes
for a hydrophobic polypeptide, i.e., chromogranin A (ChrA), have been identified
in Pseudomonas aeruginosa and Alcaligenes eutrophus and thought to be
responsible for the outward translocation of Cr (VI) anions (Cervantes and
Silver, 1992). There is also evidence that sulfate, which is chemically-analogous
to the Cr (VI) anion, can competitively inhibit Cr (VI) uptake in microbial systems,
leading to lower accumulation of Cr (VI) and bacterial resistance (Ohtake et a!.,
1987). Yet another mechanism of the Cr (VI) resistance has been discovered in
Pseudomonas ambigua (Horitsu et a!., 1983). The particular surface structure (a
thick envelope) of the bacterial membrane results in its low permeability to Cr
(VI). The unique abilities of bacteria to reduce and withstand toxic Cr (VI) may
be useful for bioremediation, an emerging biological method to remediate Crcontaminated soils.

2.3.

Heavy Metals and Plants

2.3.1. Metal toxicity

Plants require both macronutrients and micronutrients for their growth.
Some trace elements (heavy metals) including Cu, Mn, and Fe are important in
plant physiological processes (Baker and Brooks, 1989), where they can act as

14

activators of a number of enzymes in photosynthesis, respiration, oxidative
phosphorylation, and DNA, ribonucleic acid (RNA) and protein syntheses
(Salisbury and Ross, 1992). However, other metals such as Co, Ni, and Al, are
not considered to have any essential function in plants (Baker and Brooks,
1989).

Whether essential or non-essential for plant nutrition, heavy metals may
become toxic to plants at relatively low concentrations. For example, Rhoads
(1971) observed toxicity in Nicotiana tabacum (tobacco) irrigated with water
containing greater than 1.5 mg Fe kg'^ soil. Excessive heavy metal ions may
induce a series of biochemical and physiological effects such as membrane
damage, alteration of enzyme activity and hormone balance, deficiency of
essential nutrients and water, and inhibition of photosynthesis and root growth
(Barcelo and Porshenrieder, 1990). The most commonly observed symptoms of
phytotoxicity are chlorosis and stunting. The chlorosis from excess of heavy
metals such as Zn, Cu, Ni, and Cd has often been attributed to failure in Fe
metabolism of treated plants (Foy et a i, 1978). Iron deficiency in plants results in
inhibition

of

both

chloroplast

development

and

chlorophyll

biosynthesis

(Imsande, 1998).

The stunting effects of metals can be commonly due to either a specific
toxicity of a metal to a plant or inhibition of root penetration into the soil. In higher

15

land plants, the root is the first organ to contact the metal, so that toxicity is first
experienced there. Inhibition of root elongation leads to decrease in uptake of
nutrients and, consequently, in plant growth. Some of the factors found to be
responsible for decreased root growth are inhibition of root cell division by Al
(Taylor, 1988), inhibition of root cell elongation by Cu and Zn (Wainwright and
Woolhouse, 1977), and the extension of the cell cycle by Zn (Povell et al., 1986).
Metal toxicity can affect not only the length of the primary root, but also the
morphology of the whole root system. In particular, toxic concentrations of Al,
Pb, Cd, and Mn have been reported to enhance lateral root formation, thereby
making the root system denser and more compact (Breckle, 1989). In contrast,
the root hair density is generally decreased. These morphological changes may
alter the root hormone balance and lower the capacity of plants to explore the
soil for water and nutrients (Barcelo and Porschenrieder,

1990). Another

consequence of high metal availability is browning of tap root tips and lateral
roots (Foy at a!., 1978). The browning appears to be due to metal-stimulated
suberization.

Root

lignification

is also

enhanced

(Paivoke,

1983).

Both

suberization and lignification of roots may limit water uptake and membrane
water permeability of plant cells (Breckle, 1989). This could result in a decrease
of the diameter of xylem elements and water conductivity in metal-exposed
plants. For example, toxic levels of Cd and Al have decreased both vessel
diameter and number of vessels in Phaseolus vulgaris (bush bean) and Zea
mays (maize) (Bennet at al., 1985; Barcelo at al., 1988). As a result of

16

insufficient absorption of nutrients and water from metal-damaged roots, shoot
cell enlargement is also reduced. In a study of Aidid and Okamoto (1993),
elongation growth rate of stem cells of Impatiens balsamina (balsam) was
inhibited by Pb, Cd and Zn due to their suppression on both cell turgor and cell
wall extensibility.

2.3.2. Mechanisms of metal uptake

One of the most important factors affecting plant accumulation of metals
is their mobility and, thus, availability in soil. High concentration of heavy metals
in a soil solution does not necessarily imply their high availability for plant
uptake. Although it is generally assumed that plants prefer the free or
uncompleted metal ion, there are a number of soil factors controlling metal
availability.

Some of the factors include presence of organic matter (quantity

and quality: particulate and dissolved), hydrous ion oxides, Al hydroxides, clay
minerals, carbonates,

phosphates, sulfates, and silicates. Adsorption and

precipitation are the main reactions which are both time and pH dependent and
somewhat governed by ionic strength (McBride, 1994). For example, while some
metals, such as Zn and Cd, are mostly present in a soluble or exchangeable
and, thus, readily bioavailable form, others, such as Pb, occur as insoluble
precipitates (phosphates, carbonates, and hydroxides), which are unavailable for
plant uptake (Pitchel et a/., 1999). However, plants have developed specific
mechanisms to solubilize metals in soil, thereby enhancing their extraction.

17

Some plants release phytosiderophores (for example, mugenic and avenic
acids) under Cu, Zn, Mn, and Fe deficiencies (Marschner, 1986; Romheld,
1991),

whereas

others

produce

thiol-containing

metal

binding

proteins

(metallothioneins) or phytochelatins (Khan et al., 2000). Glutathione, being the
most abundant metal-binding peptide, has been observed to be induced in Cdexposed plants (Guo and Marschner, 1995; Salt at al., 1998). In addition, plants
can reduce soil-bound metal ions, including Cu and Fe, by specific plasma
membrane-bound enzymes (metal reductases) (Salt at al., 1998). Moreover,
they can release As, Cu and Mn, but immobilize Cr by exchange with protons
(Lepp, 1981). Finally, low-molecular-weight organic acids, originating from root
exudates (for example, citric, oxalic, and malic acid), can also complex Cu, Pb,
and Cd in the plant vacuole (Marschner, 1986). It should be noted that all of the
above-mentioned processes of metal solubilization could also be performed by
rhizospheric microorganisms, such as mycorrhizal fungi or root-colonizing
bacteria (Salt at al., 1998). For example, Davies at al. (2001) reported a 3-fold
increase of Cr (III) accumulation in the roots and a 3.4-fold increase of Cr (VI)
level in the leaves of Halianthus annuus (sunflower) with an arbuscular
mycorrhiza genus Glomus intraradicas. Further, Salt at al. (1995) illustrated 3fold increase of Cd concentration (0.1 mg L'^ exposed) in the shoots of 2 weekold B. juncaa seedlings with root-colonizing strains of Pseudomonas putida and
Bacillus thuringiansis.

18

Once in the soil solution, metal ions can enter plant root cells either via
cell walls (apoplastically) or the plasmodesmata (symplastically). The electrical
charge of metal ions prevents them from moving freely across the lipophilic
cellular membranes (Lasat, 2002). Therefore, most metal ions enter plant cells
by an energy-dependent process via specific channels or ion carriers (transport
proteins) (Clarkson and Luttge, 1989), while metal-chelated complexes can be
transported via specialized carriers (Crowley et al., 1991).

2.4.

Brassies juncea

2.4.1. Geographical distribution

Brassies juneea is of Himalayan origin and is grown widely in the north
portion of the Indian subcontinent and in various parts of China (Kimber and
McGregor, 1995). In addition, it has been grown in the temperate regions of the
world, particularly on the American Great Plains, in Hungary, and in Britain
(Encyclopedia Britannica, 2003). In Canada, this crop shows high yield in the
southern low-rainfall prairie areas (Mendham and Salisbury, 1995).

2.4.2. Use

The crops in the mustard family are principally used as spices. However,
because of their unpalatable flavor, the seeds of B. juneea are unsuitable for

19

condiment purposes. Therefore, the B. juncea cultivars are mostly grown for
crushing to obtain oil (Tsunoda e tal., 1980).

2.4.3. Life cycle

S. juncea is a fast-growing (90 to 95 days from sowing), cool-season
(early

spring)

annual

crop

and

seeds

are

harvested

in early

autumn

(Encyclopedia Britannica, 2003). All mustards, including B. juncea, have four
distinct phases of development. During seedling phase, the above-ground mass
of the plants consists of the hypocotyl and two photosynthetic cotyledons, which
in B. juncea are wide with deep notches (Tsunoda et al., 1980). The seedling
phase lasts from 7 to 10 days. The next developmental phase of mustards is
vegetative growth, in which plants produce leaves for 3 to 6 weeks. B. juncea
has simple pinnate leaves with alternate arrangement (Tsunoda e ta l., 1980) and
stems that are glaucous to bluish green in color (Hemingway, 1995). From the
vegetative growth stage, the plants grow rapidly and enter a phase of dense
flowering. Flowering is indeterminate and begins from the base to the top of the
inflorescence. At flowering, plants bolt, opening 4 to 5 yellow flowers per day
over 2 to 3 weeks. The flowers remain open for about 3-4 days before final
shedding of petals (Labana and Suringer, 1984). Two of the six stamens in the
flower structure are lower and shorter than others (Encyclopedia Britannica,
2003). Flowers of B. juncea are easily pollinated by wind and insects, however,
they are about 80% self-pollinating (Labana and Suringer, 1984). The final

20

growth stage of mustards is fruiting. It lasts 6 to 10 weeks after sowing, and it
terminates by the senescence of the sickle-shaped green pods. The seed pod
usually consists of two outside walls, separated by a thin white partition
(Encyclopedia Britannica, 2003). The pods of B. Juncea contain up to 20 seeds
each and the crop averages 408,000 seeds kg'^ (Hemingway, 1995). The
mature plant can reach a maximum height of 4 feet (USDA NRCS, 2003).

2.4.4. Growth requirements

2.4.4.1.

Soil type

B. Juncea has many valuable characteristics that make it unique among
the other members of the mustard family. It can easily grow on many different
types of soils. For example, this species is well adapted to fine (clay) and
medium-textured (loam) soils (Madson, 1951). Although B. Juncea prefers
neutral to slightly alkaline soil conditions, it can tolerate and grow normally at pH
as low as 5.6 (Hemingway, 1995).

2.4.4.2.

Soil fertility

B. Juncea does not require more than normal soil levels of nitrogen (N),
potassium (K) or phosphorous (P).However,
(60, 40, and 20 kg ha'^ N,

P 2 O 5 , and K2 O,

in this plant,improved fertilization
respectively)

hassignificantly

increased its growth (plant height and leaf area) and seed yield when irrigated

21

with saline waters of different concentrations (50, 100, and 150 meq L'^) (Garg
etal., 1993). These authors suggested that an improvement in the concentration
and uptake of N, P, and K under high fertility stimulates the activity of nitrate
reductase and other ammonia assimilating enzymes. This causes the level of
soluble protein and amino acids to increase greatly and contribute to the
improved plant performance under salt stress.

2.4.4.3.

Temperature

Studies have shown that B. juncea can tolerate spring frosts without
serious harm (Hemingway, 1995). In contrast, this plant has shown to be
sensitive to heat stress. For example, Angadi e ta l. (2000) investigated the effect
of short periods of high temperature stress on the reproductive development and
yield of Brassica species. The researchers found that high temperature
treatment,

i.e.,

35°C,

was

harmful

to

reproductive

organs

at

different

developmental stages. In particular, it caused a decrease both in the number of
seeds produced by the main stem and the number of fertile pods.

2.4.4.4.

Water

B. Juncea can be easily grown on land of adequate moisture supply. It is
more tolerant of drought than Brassica napus (canola) but less than Triticum

22

aestivum (wheat). Good moisture prolongs flowering, which results in increased
seed number and higher yield (Hemingway, 1995).

2.4.5. Heavy metal uptake

B. juncea has the remarkable capacity to accumulate high levels of heavy
metals. For example, its shoots have been reported to take up as high as
308,600 mg Ni kg'^ plant (DW) and 172,300 mg Zn kg'^ plant (DW ) in a fertilized
sand/Perlite mixture supplemented with 100 mg Me kg'^ soil (Kumar et al.,
1995). B. Juncea has also demonstrated a strong accumulation of strontium
radioactive isotope (^°Sr), found in the soils in the Chernobyl regions of Ukraine
(Salt et a!., 1995). In particular, the final concentration of this radionuclide in
shoots of the plant was 12-fold higher than in the soil. In another experiment, the
shoot Cu concentration in this plant was 31.2 mg kg"^ plant (DW), more than
three times greater than the mean Cu concentration in the shoots of the non­
treated plants, i.e., 10 mg kg'^ plant (DW) (Ebbs and Kochian, 1997). In addition,
the roots treated with 10"^ M Pb accumulated a substantial amount of Pb
(15,982.8 pg g'^ plant DW), which was about 184 fold of control plants (Liu et ai.,
2000). The authors have also reported that over 95% of the Pb accumulated in
treated plants was found in the roots.

23

2.4.6. Heavy metal phytotoxicity

2.4.6.1.

Effects on shoots

Kumar at al. (1995) reported that B. juncea showed mild chlorosis when
grown for 14 days in a Zn-contaminated soil. However, there was no evidence of
any toxicity of the plant grown on the same medium with 10 mg Cu kg'^ soil. The
leaves had little decrease in chlorophyll when grown in the presence of 6.5 mg
L'^ Zn and 0.32 mg L'^ Cu (Ebbs and Kochian, 1997). The same effect on
chlorophyll content has been observed in B. Juncea treated with 4mM Pb. In
contrast, no reduction in chlorophyll content has been noticed in plants treated
with 2 mM Ni (Burd et a!., 2000). Liu et al. (2000) have reported the reduction in
the number of leaves in B. juncea grown on a Pb-treated soil. In particular, while
control seedlings had 4 or 5 leaves, seedlings supplied with 10'®, 10"'^ and 10'^ M
Pb had 3, 4 and 2 leaves, respectively. Similarly, Haag-Kerwer et al. (1999)
have demonstrated inhibition in leaf expansion (the 3’^'“ and 4*^ leaves) in B.
juncea exposed to 25 pM CdNOs The same researchers have also observed the
decline in transpiration rate, while the CO 2 assimilation rate (photosynthesis) of
the plants had remained unaffected. Further, Begonia et al. (1998) found that the
total leaf area of B. juncea treated with 250 and 500 mg L'^ Pb was reduced
25% and 47% , respectively, compared to the untreated plants. The same
authors and Daniels-Davis (1996) have noted anthocyanin pigmentation or
purplish coloration of leaves of S. juncea treated with 500 mg L'^ of Pb.

24

Some researchers found a significant decrease in shoot dry weight of B.
juncea when grown on soils contaminated with heavy metals such as Pb (500
mg Pb kg'^ soil: Kumar et al., 1995; Daniels-Davis, 1996; 10'^ M Pb: Liu et a!.,
2000), and Cu (0.32 mg Cu L '\

Ebbs and Kochian, 1997), while other

researchers did not observe any effect of Pb (100, 250 and 500 mg Pb L '\
Begonia et al., 1998; 4mM Pb: Burd et al., 2000), Zn (6.5 mg Zn L '\ Ebbs and
Kochian, 1997), Cu and Cd (10 mg Cu kg'^ and 2 mg Cd k g '\ Kumar et al.,
1995) on shoot biomass of the plant. While the presence of 5 and 10 mM Zn
inhibited seedling growth, Zn at 0.05 mM promoted growth of B. Juncea
seedlings, expressed in a higher shoot length of the treated plants than that of
the control plants (Prasad e ta l., 1999).

2.4.6.2.

Effects on roots

Stunting or reduced root biomass is a commonly observed growth
response of heavy metal-treated B. juncea. For instance. Ebbs and Kochian
(1997) have reported a greater reduction in root dry weight in plants treated with
Cu compared to those treated with Zn. In the same experiment, Cu was also
shown to inhibit lateral root development. In particular, the density of lateral roots
in the Cu-treated plants was significantly less than that of the control plants. In
addition, a decrease in root length of B. juncea was observed (Liu et al., 2000).
Aside from stunting, a purplish color was noted on main roots in Pb-treated B.
juncea plants in contrast to white roots of untreated plants (Begonia et al., 1998).

25

In Cu-treated plants, primary laterals have appeared discolored with a red-brown
coloration of root tips (Ebbs and Kochian, 1997).

2.5.

Chromium and Plants

2.5.1. Chromium impact on plant growth and development

2.5.1.1.

Beneficial effects

Chromium is not considered an essential element for plants (Tinker,
1981). Nevertheless, there are several reports on stimulation of plant growth by
small concentrations of Cr. Bertrand and de Wolf (1965) have illustrated that
applications of 40 g Cr ha'^ considerably increased yields of Pisum sativum
(pea) and Daucus carota (carrot). In another study, in Russia, the addition of
K2 Cr0 4 (600 g ha'^) has been observed to improve the weight, size, sugar
content, and yield of Vitus vinifera (grape) (Dobrolyubskii and Slavvo, 1958).
Similarly, Pratt (1966) has reported that applications of K2 Cr2 0 r at 30 and 100 g
kg'^ soil increased the yield of Cucumis sativus (cucumber). The reasons for the
beneficial effect of a low Cr concentration on plants are not known. Some
theories revolve around indirect effects of Cr on mineral nutrition, antifungal
agents (Pratt, 1966), and water relations (Barcelo etal., 1986).

26

2.5.1.2.

Visible deleterious effects

The phytotoxic properties of Cr have been demonstrated in solution
culture at concentrations as low as 1-2 mg L'^ (Baker and Brooks, 1989).
Chromium is known to cause chlorosis and necrosis of leaves (Barcelo et al.,
1986) and to inhibit photosynthesis (Sharma et al., 1995). These toxicity effects
are largely concentration-dependent. For example, in Avena sativum (oat), when
supplied with 5 and 10 mg Cr (VI), as K2 Cr 0 4 , kg"^ soil, the effect was one of
chlorosis, while, in the same plants, supplied with 15, 25, and 50 mg Cr (VI) kg'^
soil, specific symptoms of Cr toxicity appeared, including plant stunting and
brownish-red coloration of leaves (Hunter and Vergnano, 1953). Gupta et al.
(1994) found a decline in total chlorophyll content in a dose-dependent manner
both in Bacopa monnieri (bacopa) and Scirpus lacustris (bulrush). Reduction in
chlorophyll content in upper leaves of Cr-treated plants is thought to be due to
inhibition of Fe and translocation and stimulation of Fe-deficiency symptoms
(Barcelo et al.,

1985). The growth reduction and chlorosis may also be

considered as consequences of toxic Cr effects in roots principally caused by
alterations in the content of essential mineral nutrients (Barcelo et al., 1985).
Inhibition of root elongation has been observed in seedlings of P. vulgaris,
treated with 96 pM Cr (VI) for 21 days (Vazquez et al., 1987), and Salvia sclarea
(cedar sage), treated with 17-34 pM Cr (VI) for 48 hours (Corradi et al., 1993).

27

2 .5 .1.3.

Cr (III) and Cr (VI) phytotoxicities

The prevailing view is that Cr (VI) is more toxic to plants than Cr (III). For
example, Shahandeh and Hossner (2000) demonstrated that Cr (VI) application
decreased the index of tolerance for B. juncea plants more than Cr (III)
application, although it was noted that the concentration of the former in roots
and shoots was greater than that of the latter. In another experiment, although
both Cr (III) and Cr (VI) inhibited both root and shoot growth characteristics of T.
aestivum, the most pronounced symptoms were observed in the roots and with
Cr (VI) treatment (Mukhopadhyay and Aery, 2000). In particular, 65% reduction
in shoot fresh weight was observed in plants treated with 2 0 mg L'^ of K2 Cr2 0 7 ,
while about 50%

reduction was found in plants supplied with the same

concentration of CrCls. Moreover, in the Cr (Vl)-stressed plants, the length of
roots declined by 80% compared to 47% of that in the shoots (Mukhopadhyay
and Aery, 2000). The observed differences in toxicity between the two forms of
Cr can be explained by the low solubility and, hence, low bioavailability (more
difficult penetration of cell membranes) of Cr (III) in soil (Barlett and James,
1996) as well as a greater tendency of Cr (III) to form large hydroxyl-polymers
with many ligands at neutral pH level (Mukhopadhyay and Aery, 2000). On the
other hand, Cr (VI)
membranes.

is more mobile in soils and easily penetrates cell

However,

if

conditions

are

adjusted

to

achieve

equal

concentrations of Cr (III) and Cr (VI), so that both Cr forms are available to
plants, toxicity may occur. For instance, chlorotic leaves and poor plant growth

28

have been noticed in Brassica oleracea (broccoli) grown for 55 days with a
nutrient solution containing 10 mg kg’^ of Cr (III) and 1 and 10 mg kg^ of Cr (VI)
(Hara and Sonoda, 1979). Similarly, McGrath (1982) found that the growth of A.
sativa seedlings was nearly ceased or inhibited when supplied with 200 pM of Cr
(INI) or 20 pM of Cr (VI) in nutrient solution. Additionally, Yamaguchi and Aso
(1977) reported that 200 mg kg'^ of Cr (III) in soil decreased root elongation of
Oryza sativa (rice) and T. aestivum, while shoots were unaffected.

In yet

another study, B. oieracea responded to excess (500 pM) supply of Cr (III) by
developing a decrease in chlorophyll concentration and activity of the heme
enzymes, namely catalase and peroxidase (Pandey and Sharma, 2002). The
authors explained that the high affinity of Cr (III) for proteins allowed it to bind
these essential enzymes, thereby inactivating them.

2.5.1.4.

Morphological and anatomical alterations

Visible symptoms of metal toxicity stress appear to be a result of
morphological

and

anatomical

(at

both

cellular

and

ultracellular

levels)

modifications. For example, injuries of the root surface expressed by severely
damaged epidermal and cortical cells, damage of root cap, collapse of root hairs
and stomata and cotyledon hairs, and reduced amounts of chlorophyll and
carotenoids have been observed (Vazquez et ai., 1987; Corradi et ai., 1993).
The authors have assumed that, due to oxidation of cell wall and membrane
components by strongly oxidizing Cr (VI), an impaired function of the plasma

29

membrane leading to plasmolysis may account for alterations in the content of
essential mineral nutrients and water loss and explain reduced intercellular
spaces, changes within chloroplasts and the wilting of root and cotyledon hairs.
In addition, accumulation of abnormally high starch levels has occurred in
parenchyma cells of the vascular cylinder (xylem and phloem) in the upper part
of the root and in the pith of the stem, suggesting that reduced root growth due
to Cr (VI) may lower starch utilization in growth processes (Vazquez et al.,
1987). Moreover, the occurrence of ameboid plastids in Cr (Vl)-treated roots
(Vazquez at al.,

1987) suggests that Cr (VI) may inhibit normal plastid

development.

There are some reports that describe the influence of Cr (VI) toxicity on
the vascular system. For example, Cr (VI) has caused a substantial decrease of
the diameter of xylem vessels (Barcelo and Poschenrieder, 1990). In a similar
study, the sizes of both phloem and xylem cells have decreased in stems of Cr
(Vl)-treated P. vulgaris (Vazquez at al., 1987). In another study, the number of
vascular bundles has increased, while the vessel density, dimension of vessel
elements and number of fibers all have decreased greatly in the root and shoot
of S. lacustris, treated with Cr (VI) (Suseela at al., 2002).

Microscopic studies of leaves illustrate increased number of trichomes,
but reduced number of granal stacks when treated with Cr (VI) (Barcelo at al..

30

1988;

Vazquez

et

al.,

1987).

Furthermore,

stomata

closure

and

poor

development of the thylakold membrane system have been observed In Cr (VI)treated plants. These results are hypothesized to be due to an indirect effect of
Cr in leaves by induction of Fe deficiency.

2.5.2. Chromium absorption and distribution

The absorption and dynamics of Cr (III) and Cr (VI) in various plant parts
have been studied in nutrient solutions (Shewry and Peterson, 1974; Skeffington
at a!., 1976; Cary at a!., 1977; McGrath, 1982; Lytle at a!., 1998; Zayed at a/.,
1998; Arteaga at a!., 2000; Davies at a!., 2001; Aldrich at a!., 2003), artificial soil
mixtures (Parr and Taylor, 1980; Barcelo at a/., 1986; Kumar at a/., 1995;
Suseela at al., 2002), and soils (McGrath, 1982; Naqvi and Rizvi, 2000;
Shahandeh and Hossner, 2000; Singh, 2001; Fernandes at al., 2002). In
laboratory experiments, both Cr (III) and Cr (VI) accumulate mainly in roots and
are poorly translocated to shoots, although absorption and translocation of the
latter is higher than that of the former (Shewry and Peterson, 1974; Barcelo at
al., 1986; Zayed at al., 1998; Shahandeh and Hossner, 2000). A separate
uptake mechanism is believed to exist for the two chromium forms. Cr (VI) is
taken up actively by the sulfate carrier, while Cr (III) is absorbed passively
(Shewry and Peterson, 1974; Skeffington at al., 1976), being bound to the
cation-exchange sites of the cell walls (Marschner, 1986). Moreover, Cr (III),
chelated in soils, has been observed to enter plant roots at slower rates than

31

non-chelated Cr (III) and Cr (VI). Nevertheless, complexed Cr (III) moves easily
to the shoots (Verfaillie, 1974). Both Cr (III) and Cr (VI) enter the vascular tissue
with difficulty; however, once in the xylem, they move more freely (Skeffington et
al., 1976).

Uptake of Cr by plants and subsequently its accumulation in plant tissues
is influenced by the amount of added metal (Barcelo et a/., 1985). In particular,
Cr absorption has been enhanced with increasing concentration of the applied
metal (Davies et ai., 2001; Shahandeh and Hossner, 2000; Fernandes et a!.,
2002). In addition, probably due to the chemical similarity of the ions, the uptake
of Cr (VI) is most likely to be severely inhibited by the sulfate normally present in
soils (Shewry and Peterson, 1974).

According to some researchers, Cr (VI) is readily translocated within
plants (Swamy, 1996; Fernandes et ai., 2002), while others have reported an
initial reduction of Cr (VI) to mobile Cr (V) (Micera and Dessi, 1988) and to Cr
(III) species (Lytle et a/., 1998; Zayed et al., 1998; Aldrich et al., 2003). For
instance, in their studies with Hordeum vulgare (barley), Shewry and Peterson
(1974) have found most of Cr in a soluble non-particulate form. They observed
that two-thirds of internal Cr was located in vacuoles of root cells, while most of
the remaining Cr was within the cell walls. The authors concluded that Cr is
unavailable for transport mainly due to its spatial localization in a specific

32

subcellular compartment in the root cells i.e., the vacuole, or the lack of a
specific mechanism

for transport.

Sanita di Toppi et al.

(2002) further

hypothesized that Cr solubilization followed by detoxification might be performed
by chelation and compartmentalization in the vacuole by low-molecular-weight
organic acids originating from root exudates. The X-ray absorption spectroscopy
(XAS) data presented by Arteaga at al. (2000) also showed that Cr (VI)
absorbed from solution was partially reduced to Cr (III) in the roots of Larrea
tridentate (creosote bush). Some Cr (VI) and the reduced Cr (III) were further
transported through the stems and finally accumulated as Cr (III) in the leaves of
the plant, bound to oxygen-containing ligands. The form in which Cr is taken up
and translocated in plants may therefore vary in different plant species.

33

3.0.

MATERIALS AND M ETHODS

3.1.

Soil Source and Preparation
Surface horizon (0 to 30 cm) of Orthic Humo-Ferric Podzol from Aleza

Lake (Research Forests, BC, Canada) with known chemical and physical
characteristics (Arocena and Sanborn, 1999) was collected (Table 3.1, Table 1:
Appendix). Fresh-field soil sample was mixed thoroughly and sieved through a
4-mm polyethylene sieve before analysis in order to preserve some of soil crumb
structure and aeration status of field soils during storage (Bartlett and James,
1996). Half of the soil was air-dried in a laboratory and passed through a 2-mm
sieve, while the other half was stored field-moist in a refrigerator at 4°C (Fisher
Scientific, Indiana, PA, USA).

The soil was analyzed for moisture content by a gravimetric method with
oven-drying at 105°C (Yamato, Japan) for 24 h (Kaira and Maynard, 1991). In
addition, background soil total Cr concentration was estimated by using
inductively coupled plasma-atomic emission spectrometry (ICP-AES) (EPA,
1996a) following acid digestion of 0.2 g soil (EPA, 1996b). SO-2 (16 mg Cr kg'^
soil DW) and SO-4 (61 mg Cr kg'^ soil DW) Canadian certified reference soils
(Canada Centre for Mineral and Energy Technology, Ottawa, ON, Canada) were
used to ensure the accuracy of the measurements.

34

Table 3.1. Physical and chemical properties of soil from Aleza Lake.

Soil Properties

Orthic
Humo-Ferric
Podzol

Sand (weight %)*

17.9

Silt (weight %)*

67.7

Clay (< 2pm)
(weight %)*

14.3

Total C (%)*

1.38

Moisture content (0m)
(kg kg-^)

0 .1 2

pH in H 2 0 ( 1 :2 )*

5.13

CEC (cmolc g'^)*

5.01

Total Cr (mg kg'^)

55.3

Alp (%)*

0.48

Pep (%)*

0 .6 8

* From Arocena and Sanborn, 1999

3.2. Chromium Application

In order to ensure easy access to the roots of Brassica juncea, two
plexiglass-made rhizotron units, covered with aluminum foil to reduce light

35

penetration to plant roots, were used for plant growth (Figure 3.1). Prior to filling
each unit of a rhizotron with a 450 g of soil (DW), the air-dried soil samples were
brought to the same water content as the field-moist ones. Upon addition of a
controlled release mini-fertilizer N-P-K (18-5-9) (Scott Osmocote, Marysville,
OH, USA) and aqueous Cr (III) as CrCl3.6H20 and Cr (VI) as K2Cr20y standard
solutions at the rate of 100 mg Cr (III or VI) kg‘^ soil (DW), both field-moist and
air-dried soil samples were mixed thoroughly by hand and the moisture of the
soils was adjusted to field capacity. Three replicates were done for each
treatment.

Removable
Plexiglass Plate

Figure 3.1. Rhizotron for growing Brassica juncea.

36

3.3.

Plant Material

Seeds

of B. juncea

(accession

FT

182921)

were

obtained

from

USDA/ARS Plant Introduction Station of Iowa State University (USA). Six seeds
of the plant were sown in each unit of a rhizotron, previously supplied with both
fertilized and Cr-treated soil.

3.4.

Plant Growth and Harvest Conditions
The

plants

were

grown

in

the

temperature-

and

light-controlled

environment of a greenhouse (Enhanced Forestry Laboratory, UNBO, BO,
Canada) with a 16-h photoperiod and a 21/18°C day/night temperature regime.
The continuous irrigation system was built with a porous polymer tube set at
around field capacity water potential, i.e., -30 kPa (Brady and Weil, 2002).
Deionized (reverse osmosis) water was used for irrigation. The experiment was
laid out on a greenhouse bench in a blocking design (three blocks, each
representing one harvest time) with a completely randomized arrangement of
treatments in each rhizotron unit within each block. At one week after
emergence, the plants were thinned to three seedlings within each rhizotron unit.

The

plants

were

harvested

at

vegetative,

flowering,

and

fruiting

developmental stages, i.e., at 17, 36, and 69 days after sowing, respectively. At

37

these times, more than a half of the control and Cr-treated plants had 3 leaves
emerged, visible buds, and filled seed pods, respectively.

3.5.

Soil Analyses

3.5.1. Soluble Or (III) and Or (VI)

Total soluble Or and Or (VI) in the rhizosphere and bulk soil samples were
extracted at each harvest time (17, 36, and 69 days after planting). Rhizosphere
soil (3mm from the roots) was separated from bulk soil by gentle scooping. Crtreated and control soil samples (1.5 g each:

DW) were shaken on a

reciprocating shaker (Eberbach Corp., Ann-Harbor, Ml, USA) for 16 h with 15
mL of deionized water (Bolan et al., 2003). All soil extracts were centrifuged at
10,000 rpm for 10 min on a Hermie Z 382 centrifuge (Mandel Scientific Comp.
Ltd., Germany) and stored a t4 °C (W CT Canada Inc., Cambridge, ON, Canada).

Total soluble Cr was quantified by ICP-AES on a Leeman Labs PS 1000
spectrometer (Leeman

Labs

Inc.,

USA),

while

Cr (VI)

was

determined

colorimetrically at 540 nm on a Spectronic 20D+ UV spectrometer (Milton Roy
Comp.,

USA)

using a modified version of the diphenylcarbazide

(DPC)

procedure (Bartlett and James, 1996). The modification consisted of reducing
the amount of a soil aliquot to 5 mL and the amount of the DPC reagent
proportionally. Analyses were conducted in triplicates. Cr (III) was estimated

38

from the difference of measured values of total chromium and dichromate
concentrations.

3.5.2. Microbial biomass C

At each harvest time, the Cr-treated and non-treated seedlings in fieldmoist and air-dried soils were gently shaken to remove the soil from the roots.
The soil adhering to the roots was defined as the rhizosphere soil, while the
remaining soil was bulk soil. The isolation of the rhizosphere soil was
accomplished in a similar procedure as described by Priha et al. (1999). In
particular, the roots were first washed with 35 mL of deionized water in glass
tubes and sonicated mildly in a water bath (Fisher Scientific, Indiana, PA, USA)
for 2 min. The solutions were centrifuged (Mandel Scientific Comp. Ltd.,
Germany) in 50-mL Nalgene plastic vials at 3,000 rpm for 10 min and the
supernatants discarded. The roots were then immersed in another 35 mL of
deionized water for further analysis of low-molecular-weight organic acids,
whereas the vials with soils were weighed to determine their fresh weight and
kept at 4°C (W C T Canada Inc., Cambridge, ON, Canada). For the Cr-amended
and control bulk soils, 5 g of fresh soil sample was weighed into tubes with 35
mL deionized water and treated in the same manner as rhizosphere soils. After
the analyses, all vials were oven-dried at 105°C (Yamato, Japan) for 24 h to
determine dry weight of the soils (A&D Comp., Japan). The average dry weight
of rhizosphere soil was 0.25 g.

39

Microbial biomass C was determined by the chloroform fumigationextraction (CFE) method described by Priha et al. (1999). Both rhizosphere and
bulk soil samples were fumigated with direct addition of 100 pi ethanol-free
chloroform at 25°C for 24 h, whereas the non-fumigated samples were kept at
4°C (W CT Canada Inc., Cambridge, ON, Canada). After the chloroform had
been removed in a vacuum dessicator (Labconco Corp., Kansas City, MO, USA)
using a vacuum pump (Welch Vacuum, Thomas Industries Inc., Skokie, IL,
USA), both fumigated and non-fumigated samples were extracted with 8 mL of
0.5 M K2 SO 4 for 30 min on a reciprocating shaker (Eberbach Corp., Ann-Harbor,
Ml, USA). The samples were then centrifuged (Mandel Scientific Company Ltd.,
Germany) at 1,500 rpm for 10 min and the extracts stored at -20°C in order to
keep samples stable (Environmental Growth Chambers, Chagrin Falls, OH,
USA) prior to the analysis. The C content of the K2 SO 4 extracts was measured
by an EnviroTOC carbon analyzer (Automation Instruments Manufacturing Inc.,
Calgary, AB, Canada). No kec, i.e., an extractable part or fraction microbial C
after fumigation, was applied; only the extractable C flush, released by
fumigation (the difference between extractable C from fumigated and nonfumigated samples), was calculated.

40

3.5.3. Chromium spéciation in soils

X-ray absorption near-edge spectroscopy (XANES) was applied to
identify the oxidation state of Cr in amended rhizosphere and bulk soils. The
XANES data were collected at the National Synchrotron Light Source at
Brookhaven National Laboratory (NY, USA) on beam-line X18B using a silicon
(Si) (111) double crystal monochromator. The Cr spectra (edge energy of 5.989
keV) were recorded using a passivated implanted planar silicon (PIPS) detector.
Soil samples were packed in a sample holder with an X-ray transparent tape and
placed in a sample chamber at 45° to the X-ray beam. Several spectra of two
replicates were collected from 100 eV below to 100 eV above the Cr edge.
Spectra were also collected for Cr (III) and Cr (VI) standard compounds, namely
Cr (lll)-acetate, Cr (lll)-formate, Cr (lll)-trioxalate, Cr (lll)-chloride hexahydrated,
and Cr (Vl)-dichromate. Their selection was based primarily on their availability
and biological significance. Standards were analyzed in a solid state. Cr
reference foil was run simultaneously with each data set.

The WinXAS software package (Ressler, 2001) was used to analyze the
X-ray absorption spectroscopy (XAS) collected data. The

samples were

calibrated against the Cr edge using the first and the second degree derivatives
of the reference foil edge energy (5.989 keV). The background correction and
normalization

were

performed

on

a

pre-edge

and

a

post-edge

region,

respectively, using a first-degree polynomial. The XANES region (5.95 to 6.05

41

keV) was then extracted from the entire spectra. Data were also analyzed
quantitatively using linear combination (LC)-XANES fittings (Ressler, 2001) of
soil samples to those of standard Cr (III) and Cr (VI) compounds.

3.6.

Plant Analyses

3.6.1. Plant growth and chemical measurements

3.6.1.1.

Visual evaluation of Cr stress

Visual evaluations of stress in B. juncea, caused by Cr, were made at the
last harvest (69 days after plant sowing). The evaluation criteria included: dead
plant = 100% of wilted and/or chlorotic leaves; very stressed = between 80 and
100% of wilted and/or chlorotic leaves; moderate stress = between 50% and
80% of wilted and/or chlorotic leaves; initial stress = between 20 and 50% of
wilted and/or chlorotic leaves and healthy = 0% of wilted and/or chlorotic leaves.

3.6.1.2.

Plant height

At the final harvest, shoot height of the control and Cr-treated 6. Juncea
plants was measured from the soil surface to the shoot tip using a ruler.

42

3.6.1.3.

Shoot and root dry weight

At each exposure period, the plants from all treatments were harvested,
triple-rinsed with deionized water, and separated into roots and shoots. Their dry
weights were measured (Sartorius, Germany) after oven drying at 70°C
(Yamato, Japan) for 24 h (Kaira and Maynard, 1991).

3.6.1.4.

Chromium accumulation

At the final harvest, the roots and shoots of Cr (III, Vl)-treated and control
plants were triple-rinsed with deionized water, separated into roots and shoots,
which were then cut into small pieces and dried in an oven at 70°C (Yamato,
Japan) for 24 h prior to the analysis. Dried plant tissues were ground with an
agate mortar and a pestle and 0.25 g of the plant material was digested in
concentrated HNO3 and 30% H2O2 using a microwave procedure as described
by KaIra and Maynard (1991). Total Cr in the digests was estimated by a
Leeman Labs PS 1000 UV spectrometer (Leeman Labs Inc., USA). Standard
reference material (1573a tomato leaves: 1.99 mg Cr kg'^ plant DW) supplied by
the National Institute of Standards and Technology (Gaithersburg, MD, USA)
was used to ensure accuracy of measurements.

43

3.6.1.5.

Low-molecular-weight organic acids in root exudates

Root exudates of B. juncea were collected at the three sampling times.
After separation of rhizosphere soil for soil microbial biomass C, the roots of
both Cr (III, Vl)-treated and control plants were immersed in glass tubes
containing 35 mL of deionized water (Strom et al., 1994). Each tube was
covered with aluminum foil to avoid any possible increase in root exudation
caused by light. The tubes were then randomly placed on a greenhouse bench
and the plants were allowed to photosynthesize for 6 h. The light and
temperature conditions were the same as for the plant growth experiment. Root
exudates were collected for three plants in each replicate. Tubes with deionized
water, as blanks, were treated and analyzed in the similar manner as the
samples. Aqueous extracts were immediately frozen and stored at -2 0 °C
(Environmental Growth Chambers, Chagrin Falls, OH, USA) for further analysis.

Capillary electrophoresis (CE) was used to separate and consequently to
identify and quantify organic acids exuded by the plant roots. The mixed
standard solution was prepared from individual chemicals using nanopure (MilliQ) water. The analyses were performed using a Beckman (Fullerton, CA, USA)
model P/ACE™ MDQ capillary electrophoresis system equipped with a photo
diode array (PDA) detector (indirect detection operating at 233 nm) and a bare
fused silica capillary column (75 pm ID x 57 cm total length). CelixirOA™ pH 5.4
kit, containing pyridine-dicarboxyIic acid buffering system (MicroSolv Technology

44

Corp., Long Branch, NJ, USA), was used to fill the capillary. To identify organic
acids in root exudates, an electrokinetic (electromigration) injection of a sample
(10 kV for 10 sec) was applied, while a hydrostatic (pressure) injection mode of
separation (0.1 psi for 10 sec) was used for their quantification. To prepare
samples for the pressure injection, 1-mL subsamples of aqueous extracts were
concentrated in a freeze-dryer (LabConco Corp., Kansas City, MO, USA)
overnight, just prior to the day of their analysis. Accuracy was accessed by
running a mixture of standards (six times) of each acid of intermediate
concentration, i.e., 1 0 0 pmol L '\ within the same day as the samples.

3.6.2. Anatomical measurements

3.6.2.1.

Stem anatomical characteristics

At the final harvest, a 7-cm piece of a stem from each treated and non­
treated B. juncea was cut between the

and 4*^ nodes from the bottom of the

stem. The stem fragments were cross-sectioned manually with a double-edged
razor blade and stained with a 0.1% toluidine blue (TBO) solution.

Measurements of stem diameter, width of epidermis, cortex, primary and
secondary phloem and xylem, and pith were taken with a compound microscope
(Olympus, Japan). The width of each plant tissue was measured in millimeters
and the results were expressed in percent that a particular tissue occupied in the

45

plant stem. The number of xylem cells was also counted by using a compound
microscope, while number of vascular bundles was determined with a dissecting
microscope (Nikon, Japan).

3.6.2.2.

Root and leaf anatomical characteristics

At the final harvest, a 0.5-cm segment of a tap root was taken 1 cm below
the point where root and shoot joined, while a 0.5-cm segment of a lateral root
was taken 0.5 cm below the point where tap and lateral root joined. In addition, a
25-mm^ piece was cut from the center of a leaf at the 3’’'' node from the bottom of
the stem. The root and leaf segments were processed according to the general
protocol of plant preparation for microscopy (Razin, 1999). In brief, the tissues
were first fixed for 3 h with 2% glutaraldehyde in 0.05 M phosphate buffer (pH
7.2). After two rinses for 30 min and one wash overnight (at 4°C) with 0.05 M
phosphate buffer, the plant samples were post-fixed for

1

h in 1 % osmium

tetroxide in the same buffer. The tissues were then washed with 3 changes of
buffer (30 min each), dehydrated in ethanol series (25, 50, 75, 95, and 100%
ethanol for 30 min each), and embedded in a medium grade LR White resin
(Canemco

Inc.,

St.

Laurent,

QC,

Canada).

Two-micron-thick

longitudinal

sections of the roots and cross-sections of the leaves were cut using an Ultracut
E (Reichert-Jung, Austria) ultramicrotome and stained with 0.1% TBO at the
University of Alberta (Edmonton, AB, Canada).

46

The sections were viewed under a compound microscope (Olympus 3
Max, Japan) and photographed with a Cool Fix 500 (Nikon, Japan) digital
camera. All measurements were obtained using Image J software (Image
Processing and Analysis in Java Software, 2003).

For lateral and tap roots, root diameter, xylem diameter, and number and
width of large cells (greater or equal to 25 pm and 12.5 pm for tap and lateral
roots, respectively) in the entire xylem were quantified. In addition, the number
and width of all cells in a designated xylem area (200pm- and 36 pm-long for the
tap and lateral roots, respectively) were determined. To choose this xylem
region, first, the xylem diameter in each plant treatment was measured. The half
value of the smallest xylem diameter was then used as a length of a box. Cell
measurements in the tap and lateral roots were taken within this box, which was
placed in the right portion of the xylem starting from its middle (Figure 3.2). For
leaves, their thickness and thickness of palisade and spongy mesophyll were
measured in the middle part of the leaf. The number of palisade layers and veins
(vascular bundles) in the middle part of the leaf was also quantified.

3.6.2.3.

Chromium spéciation in plant tissues

Root and leaf samples of 69-day-old B. juncea, exposed to either Cr (III)
or Cr (VI), were frozen, ground to a fine powder in liquid nitrogen using a pestle
and a mortar and stored at -80°C in order to keep the plant tissues stable

47

(Sanyo, Japan). To speciate Cr, X-ray spectra of frozen plant tissues were
collected at the National Synchrotron Light Source at Brookhaven National
Laboratory (NY, USA) on beam-line X18B and analyzed with the WinXAS
software package (Ressler, 2001) in the similar manner as the soil samples (see
section 3.5.3).

1 OO u m

Figure 3.2. Longitudinally-sectioned lateral root (200x) from control Brassica
juncea showingbox used for measurements of cells in a part of xylem.

3 .6.2.4.

Chromium localization within plant

The longitudinal sections of roots and cross-sections of leaves of Cr (III,
Vl)-treated B. juncea were prepared in the same way as for light microscopy
measurements (see section 3.6.2 2).

48

The elemental distributions within roots and leaves were obtained using
synchrotron-based

X-ray

microprobe

spectroscopy.

Measurements

were

performed on beamline 20-ID at the Advanced Photon Source at Argon ne
National Laboratory (Chicago, IL, USA). Fluorescence data were collected with a
13-element germanium (Ge) detector that was placed at 45° from the specimen
and focused on a 50 pm diameter portion of the sample. The samples were
scanned in 0.01 keV energy steps (5 sec per step increments). The incident
photon energy was set at 5.989 keV for all scans.

3.7.

Statistical Analyses

Means and standard errors were calculated to summarize the data. One­
way analysis of variance (ANOVA) was performed using SPSS (Chicago, IL,
USA) for Windows statistical program (SPSS, 1989-2002) in order to compare
the means of all measured characteristics of Cr (III, Vl)-treated and control
plants. Significant differences between the means were accessed by the least
significant difference (LSD) test using the General Linear Model procedure
(SPSS Inc., 1989-2002).

49

4.0.

RESULTS AND DISCUSSION

4.1.

Soluble Cr (III) and Cr (VI) in soil

The data for water-soluble Cr (III) and Cr (VI) in field-moist and air-dried
bulk and rhizosphere soils of Brassica juncea, at 17, 36, and 69 days of growth,
are presented in Figure 4.1. The results indicate that generally low amounts of
either Cr (III) or Cr (VI) (pg kg'^ levels) were extracted from soil with water,
perhaps due to inefficiency of water as an extractant or rapid uptake of added Cr
(III, VI) compounds by the plant.

Throughout the experiment, lower amounts of soluble Cr (III) and Cr (VI)
were consistently extracted from the soils treated with 100 mg Cr (III) kg'^ soil
than from the soils amended with the same amount of Cr (VI) (Figure 4.1). The
fate of water-soluble Cr (III) compounds added to soils includes dissociation of
Cr(H 2 0 ) 6 ^^ complex to the sparingly soluble Cr (III) hydroxide and protons and
further formation of the even less soluble Cr (III) oxide (Grove and Ellis, 1980).
The formation of protons could have decreased pH of Cr (lll)-amended soils. As
a result, in the acidic Aleza Lake soil used in this study (pH = 5.13: Arocena and
Sanborn, 1999), this decrease in pH may have caused the precipitation of Cr (III)
oxide and, thus, low initial extractability of Cr (III) by water. In contrast,
dichromates are generally less likely to precipitate and are expected to be more
mobile (Kimbrough eta!., 1999).

50

1000

800
—♦—
— ■—
— ^—
—•—
—•—
— ▼—

800

600

C r (III) fie ld -m o is t rh iz o s p h e re
C r (Ml) fie ld -m o is t bulk
C r (III) air-dried rh iz o s p h e re
C r (III) air-dried buik
C r (Vi) air-dried rh izo s p h e re
C r (Vi) air-dried buik

C r (III)

600

Or (VI)
400

I
400

>
o

Ü

200
200

0

20

40

60

0

20

40

60

T i m e ( days)
Figure 4.1. Water-soluble Cr (III) and Cr Ç)/l) extracted from the Cr (III) and Cr (Vl)-amended (100 mg kg'^ of either
CrCIs 6 H 2 O or K2 Cr2 0 r) field-moist and air-dried rhizosphere and bulk soils of Brassica juncea after 17, 36, and 69
days of growth.

51

Figure 4.1 also illustrates that, during the first 17 days, about 63% of
water-extractable Cr in Cr (Vl)-treated soils was in the form of Cr (III) species
indicating the reduction of Cr (VI). Compounds with potential for reducing Cr (VI)
to Cr (III) could be soil organic matter (Bartlett and Kimble, 1976) or other
electron donors such as low-molecular-weight organic acids (Hale et al., 1978).
The latter can be formed during decomposition of organic residues in soils
derived from decaying animal and microbial tissues, or released by plant root
exudates (Hale et al., 1978). Root exudates have been reported to affect the
redox behavior of Cr because of their ability to reduce Cr (VI) and to form
soluble complexes with Cr (III) species (Lundstrom, 1993). For example, organic
acids such as citric, malic and aspartic acids released in root exudates in Z.
mays, treated with different levels of Cr (III) and Cr (VI), have been reported to
enrich Cr uptake possibly by the formation of organically-bound Cr (III)
compounds (Srivastava et a i, 1998a).

During the next 19 days of metal exposure, Cr (VI) was not detected (<
24 pg kg'^) in any soil and Cr treatments (Figure 4.1), suggesting that the
reduction of Cr (VI) to Cr (III) in the soils could have occurred. Soil pH,
bioactivity,

and

oxygen

status

are

believed

to

be

among

important

characteristics in accessing the reducing power of the soil (Losi et a i, 1994).
Organic materials (total C = 1.38%: Arocena and Sanborn, 1999) appear to be
limited in the Aleza Lake soil; therefore, plant root and microbial respiration (an

52

oxygen-consuming and carbon dioxide-releasing process) could have lowered
the O 2 level, decreased pH, and provided additional C enrichment (through root
exudation and decay), thereby favoring Cr (VI) reduction.

At the final harvest (69 days after plant sowing), in any soil and Cr
treatment, however, no water soluble Cr (III) could be detected (< 72 pg kg'^)
(Figure 4.1). Therefore, a significant amount of soluble Cr (III) may have
hydrolyzed, adsorbed and, thus, become unavailable.

As observed in Figure 4.1, after 17 days of metal exposure, in field-moist
soils treated with Cr (III), the concentration of water-extractable Cr (III) was
significantly lower, whereas the concentration of water-extractable Cr (VI) was
significantly higher in the bulk compared to the rhizosphere soil. This trend was
not observed in the case of either air-dry soils amended with Cr (III, VI) or fieldmoist soils amended with Cr (VI). Oxidation of a small portion of soluble Cr (III)
added to soils, most likely by IVInOg, could possibly have contributed to the
presence of soluble Cr (VI) in field-moist bulk soil and, thus, to the decrease of
soluble Cr (III) in the same soil. In contrast, the most obvious effects of drying
are greater solubility and reducing ability of soil organic matter (Bartlett and
James, 1988). As a result, surface acidity increases and Mn (IV) is reduced to
Mn (II), becoming exchangeable and soluble. Both lower pH and higher solubility
of organic matter probably result from the increased polarity of surface-oriented

53

water as drying proceeds (Bartlett and James, 1988). Buildup of reduced Mn (II),
attracted by the negative charge on the oxide surface, leads to slowing of the
oxidation of Cr (III), because the adsorption of reduced Mn (II) has resulted in a
plus-charged oxide surface that repels ionic Cr (III) (Bartlett, 1991). This could
explain the fact that the dried-rewet soil samples did not oxidize any Cr (III)
throughout the present study. In a similar study of Barlett and James (1979),
although only about 7% of added Cr (III) was oxidized to Cr (VI), 10- and 5-times
more Cr (VI) was formed in the moist samples than in the dried-rewet ones, after
40 hours and 15 days of Cr (III) application, respectively. On the other hand, the
absence of soluble Cr (VI) in the field-moist rhizosphere soil of Cr (lll)-treated
plants in the present experiment (Figure 4.1) demonstrates that the rhizosphere
may have prevented oxidation of Cr (III) or readily reduced formed Cr (VI) to Cr
(III).

4.2.

Chromium spéciation in soil

Information about valence state of Cr in amended soils was obtained
using XANES. XANES spectra of the reference compounds of Cr (VI) as
potassium dichromate and Cr (III) as Cr (lll)-chloride hexahyd rated, Cr (III)trioxalate, Cr (lll)-acetate, and Cr (lll)-formate are shown in Figure 4.2. As seen
from this figure, the spectra of Cr (VI) and Cr (III) compounds are very different.

54

1. 5

C r ( V l ) - d i c h r o m ate

3

S

c
0

1. 0

1
o

C r (lll)-trioxalate

È

<

■a

C r (Ul ) - f or m ate

N
(0

g
O

0.5

Cr (lll)-acetate
Cr (lU)-chloride hexahydrated^
5,95

5,96

5,97

5,98

5,99

6,00

6, 01

6,02

6,03

6,04

6,05

Photon energy (keV)
Figure 4.2. X-ray absorption near-edge spectroscopy (XANES) spectra of Cr (III, VI) reference compounds.
55

In particular, the pre-edge peak in Cr (VI) spectrum is intense but it is small and
indistinct in all Cr (III) species. In all soil and Cr treatments, the appearance of
the XANES of the rhizosphere and bulk soil samples (Figures 4.3 and 4.4) is
similar to that of model Cr (III) species (Figure 4.2). It is evident, therefore, that
Cr (III) is the valence state of Cr in all soils. These results are in agreement with
the data on water-soluble Cr (VI), which was not detected in any soil treatments
at the final plant harvest (Figure 4.1). Therefore, the reduction of Cr (VI) to Cr
(III), perhaps mediated by soil organic matter or products of its oxidative
degradation (non-humic organic acids) in the bulk soils and products of plant
and/or microbial metabolism (exudates) in the rhizosphere soils, seems to be the
major mechanism of Cr (VI) removal from soils.

The quantitative data for the linear combination (LC)-XANES fittings
(Table 4.1) further support the above conclusion. None of the soils appeared to
contain any potassium dichromate. In contrast, from the reference compounds
used in this study, Cr (lll)-formate and Cr (lll)-acetate were the best fits for the
bulk and rhizosphere soils, respectively, with the exception of Cr (lll)-trioxalate
(74%) being predominant in the Cr (lll)-amended air-dried rhizophere soil (Table
4.1).

These

findings

suggest

that

low-molecular-weight

organic

acids,

particularly formic and acetic acids, could have contributed to Cr (VI) reduction.

56

5

3

«
C

O
0

0
È

<

1
■o

te

Ë

O

0.

5.95

5 .96

5.97

5.98

5.99

6.00

6.0 1

6.02

6.03

6 .04

6.05

Photon energy (keV)
Figure 4.3. X-ray absorption near-edge spectroscopy (XANES) spectra of field-moist: (a, b) Cr (lll)-amended
rhizosphere and bulk soils; (c, d) Cr (Vl)-amended rhizosphere and bulk soils. Dashed curves represent linear
combination (LC)-XANES data fittings.
57

5

3

5

c
o

0

0

0}

<

1
■a

N.

co

Ë
O

0.5

z

5.95

5.96

5.97

5.98

5.99

6.00

6.01

6.02

6.03

6.04

6.05

Photon en e rg y (keV)
Figure 4.4. X-ray absorption near-edge spectroscopy (XANES) spectra of air-dried: (a, b) Cr (lll)-amended
rhizosphere and bulk soils; (c, d) Cr (Vl)-amended rhizosphere and bulk soils. Dashed curves represent linear
combination (LC)-XANES data fittings.
58

Table 4.1. Distribution of chromium compounds (%) in the rhizosphere and bulk field-moist and air-dried soils of
Brassica juncea treated with 100 mg kg'^ of either CrCIs OHgO or K2 Cr2 Ü 7 (n = 2).

Cr (Vi)-amended soii

Cr (lil)-amended soii
Reference
compound

Field-moist

Fieid-moist

Air-dried

Air-dried

Rhizo­
sphere

Buik

Rhizo­
sphere

Buik

Rhizo­
sphere

Buik

Rhizo­
sphere

Buik

C r (Vl)-dichromate

0

0

0

0

0

0

0.0

0

C r (lll)-chloride
hexahydrated

2

0

0

0

0

0

10

17

Cr (lll)-formate

19

98

0

66

0

70

15

78

Cr (lll)-acetate

79

2

26

18

79

30

75

5

Cr (lll)-trioxalate

0

0

74

16

21

0

0

0

59

Being the simplest organic acid, formic acid in plants can be a product of
photorespiration, fermentation, and possibly direct CO 2 reduction in chloroplasts
(Igamberdiev et al., 1999). In addition, it can be produced during soil organic
matter decomposition (Stevenson, 1967). Moreover, soil bacteria may produce
formic acid by utilizing oxalate as a C source in a series of reactions involving
coenzyme A (CoA) (Hodgkinson, 1977). Acetic acid can also be formed in higher
plants in an irreversible reaction of pyruvic decarboxylation metabolic pathway
and during microbial fermentation (Robinson, 1986). Most low-molecular-weight
organic acids are water-soluble. They can release protons and the anionic forms
can function as ligands, which through surface and in-solution complexation
reactions affect metals solubility and spéciation (Harter and Naidu,

1995).

Monovalent organic acids, including formic and acetic acids, are weakly adsorbed
to the soil’s solid phase (Jones et al., 2003), thereby making them potential
chelates of reduced Cr (III).

4.3.

Chromium influence on soil microbial activity

The values of the C flush for the Cr (III, Vl)-treated and control field-moist
and air-dried soils are presented in Table 4.2. The C flush was generally higher
(p < 0.05) for the rhizosphere than for the bulk soil in all treatments. To quantify
the rhizosphere effect, an R/E, i.e., rhizosphere over edaphosphere or bulk soil,
ratio can be used. The R/E ratio is determined by dividing the activity of

60

Table 4.2. Chloroform fumigation-extraction C flush (mg C kg'^ DW soil) in field-moist and air-dried bulk and
rhizosphere soils of Brassica juncea treated with 100 mg kg'^ of either GrCla.OHaO or K2 CÏ2 O 7 .
17 days

Field-moist
Treatment

Rhizo­
sphere

Control

22"
(1.1)'
22"
(0.3)
18"
(0.6)

5.22
(0.049)^

Cr(lll)
Cr (VI)

treatment

36 days

Air-dry

Fieid-moist

Bulk

Rhizo­
sphere

3°
(0.6)
3"
(0.3)
0.7"^
(0.08)

22"
(1.2)
22"
(0.3)
17"
(0.6)

(0.08)
0.5"
(0.11)

5.53
(0.044)

5.94
(0.038)

12.52
(0.007)

69 days

Air-dry

Fieid-moist

Air-dry

Buik

Rhizo­
sphere

Bulk

Rhizo­
sphere

Bulk

Rhizo­
sphere

Bulk

Rhizo­
sphere

2°

44"
(2)
42"
(1.3)
35"
(1.2)

4"
(1.2)
4"
(0.13)
3"
(0.2)

44"
(2)
41"
(0.4)
34"
(2)

4"
(0.11)
4"
(0.3)
2"
(0.2)

57"
(0.5)
57"
45"
(2)

6"
(0.2)
6"
(0.6)
4'*
(0.3)

57"
(2)
56"
(2)
46"
(0.5)

(0.4)
6"
(0.7)
3“
(0.2)

6.48
(0.032)

5.24
(0.048)

14.74
(0.005)

5.31
(0.047)

(0.2)
2b

Treatment

Fieid-moist

Air-dry

Control
C r(lll)
C r(V I)

175.19(0.000)
1142.33 (0.000)
651.23 (0.000)

180.52 (0.000)
2132.02 (0.000)
536.08 (0.000)

Treatment
Control
C r(lll)
C r(V I)

Rhizosphere
0.00(0.991)
0.27(0.631)
0.60 (0.483)

Buik
1.01 (0.372)
0.08(0.851)
1.24 (0.327)

F ratio and (p value)
22.76
14.83
7.74
9.18
(0.022)
(0.002)
(0.015)
(0.005)
rhizosphere x bulk
Fieid-moist
Air-dry
441.71 (0.000)
430.27 (0.000)
4165.47(0.000)
560.42 (0.000)
449.98 (0.000)
237.88 (0.000)
field-m oist x air-dry
Buik
Rhizosphere
0.04 (0.843)
1.12(0.350)
0.27(0.631)
1.70(0.262)
3.78(0.124)
0.22 (0.666)

(3)

Bulk
6°

Fieid-moist

Air-dry

5849.61 (0.000)
191.02 (0.000)
237.30 (0.000)

674.73 (0.000)
646.43 (0.000)
4858.00 (0.000)

Rhizosphere
0.01 (0.945)
0.07(0.801)
0.09(0.780)

Buik
0.68(0.457)
0.11 (0.756)
2.86(0.166)

^ Standard errors. ^ p value. No kec correction. Within each time, means followed by a common letter are not significantly different from
each other using one-way ANOVA and LSD test (a = 0.05, n = 3).

61

microorganisms (C flush in a gram of the rhizosphere soil) by the C flush in a
gram of the bulk soil. In the present study, the R/E ratios varied from 8.32 to
34.92. In the rhizosphere, there is a continuous flow of organic substrates
derived from roots including exudates, leaked and secreted chemicals, sloughed
root cells,

and

mucilages

which

can

be

readily

used

as

nutrients

by

microorganisms (Wardle, 1992; Curl and Truelove, 1996). As a result, microbial
biomass and activity are generally higher in rhizosphere than in bulk soil. Jensen
and Sorensen (1994) also found that in the H. vulgare rhizosphere, the SIR
(substrate-induced respiration) rates were 72-170% higher than those in the bulk
soil. In another study, Priha et al. (1999) observed increase in flushes of 0 and
N in rhizosphere soils of Pinus sylvestris

(Scots pine),

Picea abies (Norway

spruce) and Betula pendula (silver birch) as compared to the flushes measured
in bulk soils.

At any Or exposure period, in both rhizosphere and bulk soils, there was
a significant decrease in the 0 flush due to Or (VI) contamination, while there
was no significant change (p > 0.05) observed in the soils treated with Or (III)
compared to those of the control (Table 4.2). Moreover, the extent of Or (VI)
inhibition did not decline with time, indicating that a permanent damage of
microorganisms

might have occurred.

Or (VI)

appeared

to be toxic to

microorganisms perhaps due to its higher availability and hence, biological
activity in soil (Ross et a!., 1981; Bartlett and James, 1988). Reports on Or

62

toxicity to soil microorganisms indicate that Cr (III) is not considered to be
particularly harmful (Ross et al., 1981; Doelman and Haanstra, 1984, 1986;
Yadav et a!., 1986), while Cr (VI) is shown to strongly inhibit most of biological
properties such as enzyme activities, basal respiration, microbial biomass 0 ,
and denitrification (Ross et a!., 1981; Speir et a!., 1995). Despite the fact that
water-extractable Cr (VI) declined markedly with time (Figure 4.1), the long-term
inhibition of microorganisms caused by this form of Cr could possibly be due to a
permanent damage of soil microbial population (Ross eta!., 1981).

Table 4.2 also shows that, during any period of Cr exposure, the
estimates of C flush decreased in the bulk soils which were air-dried and then
rewetted prior to Cr application compared to those which were used as fieldmoist. However, these changes were not significantly different (p > 0.05) in any
treatments.

Although

microorganisms

can

be

killed

during

desiccation

(Sorensen, 1983), 70-87% of microbial biomass may eventually recover after
soils are remoistened mainly due to the decomposition of various sources of
organic matter that the air-drying process made available (Shan-Min et a!.,
1987). In this study, on average, the C flush recovery was 81% (Table 4.2).

63

4.4.

Macroscopic effects of chromium on plant growth

4.4.1. Chromium uptake

Chromium concentration in shoots and roots of 6. Juncea in the fieldmoist and air-dried soils, treated with either Cr (III) or Cr (VI), is shown in Table
4.3. There were no significant differences (p < 0.05) in either root or shoot
concentration in Cr (lll)-treated plants grown in field-moist and air-dried soils.
This could be due to the fact that only a small portion of Cr (III) oxidized to Cr
(VI) (Figure 4.1), thereby not affecting Cr concentration in plant tissues (Table
4.3).

In all soils, Cr concentration in the roots was 18-40 times higher than in
the shoots (Table 4.3). In addition, the plants accumulated more Cr from the
field-moist or air-dried soils supplied with Cr (VI) than those treated with Cr (III).
The Cr concentration in the plant tissues, however, does not take plant biomass
into consideration and, thus, might not be an accurate evaluation of the ability of
B. juncea to extract Cr from the soil. Therefore, the total amount of Cr taken up
by the roots and transported to the plant shoots was also calculated (Table 4.4).
If the plant biomass is taken into account, during the growth period, the Cr (III)treated plants, grown in the field-moist and air-dried soils, removed an average
of 49 and 47.7 pg Cr per plant (148 and 143 pg or 0.3% per rhizotron).

64

Table 4.3. Chromium concentration in roots and shoots of Brassies Juncea after 69 days of growth in Cr (III, VI)contaminated field-moist and air-dried soils.

Treatment

Roots
(mg C rkg''' DW)

Control moist

BDL^

Control dry

BCF^

Shoots
(mg C rkg''' DW)

Root/Shoot
ratio

Index o f tolerance^

BCF

2.4»
(0.53)
2.9»
(0.32)

BDL*

Cr (III) moist

208^
(5.5)

2.1

5.9f
(0.16)

0.059

35

88

Cr (III) dry

20Qb
(5.1)4

2.0

5.6'=
(0.33)

0.056

36

87

Cr (VI) moist

390=
(7.1)

3.9

20=
(1.05)

0.20

19

70

Cr (VI) dry

388=
(7 6)
34.57(0.000)^

3.9

21 =
(1.43)
81.84 (0.000)

0.21

18

68

-

-

-

F ratio and
(p value)

-

' Bioconcentration factor = Cr (ill, VI) concentration in plant tissue (mg kg'^) at harvest / concentration of added Cr (III, VI) in soil (mg kg‘^).
^ Index of tolerance (%) = (total DW of Cr (III), (Vl)-treated plant / total DW of control plant) x 100.
^ Below detection limit (< 4.5 pg Cr kg'^ DW plant).
^ Standard error. ® p value. Within each column, means followed by a common letter are not significantly different from each other using
one-way ANOVA and LSD test (a = 0.05, n = 3).

65

Table 4.4. Chromium accumulation^ in roots and shoots of Brassies juncea after
69 days of growth in Cr (III, VI)-contaminated field-moist and air-dried soils.

Treatment

Root uptake
(/xg Crplant'^ )

Shoot uptake
(jug Crplant'^ )

Total root and shoot
uptake
(jug C r rhizotron^)

Control m oist

BDL^

2.2*
(0.48)

6.6

BDL*

2.7*
(0.29)

8.0

C r (III) m oist

45"
(1.4)3

4 .0 “’
(0.13)

148

C r (III) dry

44b

3.7“’
(0.21)

143

(1.3)
C r (VI) moist

47 “’
(0.9)

10==
(0.52)

173

C r (VI) dry

48 “’
(1.0)

I f
(0.77)

180

378.74

49.60
(0.000)

Control dry

F ratio and
(p value)

(0 .0 0 0 /

-

^Root (shoot) concentration (|jg g'^) x root (shoot) DW (g).
^Below detection limit (< 4.5 |xg Cr kg'^ DW plant).
^ Standard error. ^ p value.
Within each column, means followed by a common letter are not significantly different from each
other using one-way ANOVA and LSD test (a = 0.05, n = 3).

66

respectively, while Cr (Vl)-treated plants, grown in the same soil treatments,
removed 57 and 59 pg Cr per plant (173 and 180 pg or 0.4% per rhizotron).
Although the Cr (Vl)-treated plants had a higher Cr concentration in their roots
(Table 4.3), the Cr (lll)-treated plants accumulated the similar amount of Cr
(Table 4.4), due to their greater biomass (Figure 4.6). On the contrary, the
values for the shoot accumulation were significantly higher for the Cr (Vl)-treated
plants compared to those of the Cr (lll)-treated plants, thereby confirming
greater Cr translocation in the Cr (VI) treatment. The tendency of Cr (III) to be
retained on the cation-exchange sites of the root cell walls (Marschner, 1986)
might explain why this Cr form was less available to the plant shoots. The
increased translocation in Cr (Vl)-treated B. Juncea observed in this study might
be due to damaged plant root membranes and transport of this form of Cr by
simple diffusion. These results are in agreement with previous studies on Cr
accumulation in diverse plant species (Cary et al., 1977; W allace et al., 1977;
Parr and Taylor, 1980; Shahandeh and Hossner, 2000; Davies et al., 2001).

The root to shoot Cr concentration ratio was 35 or 36 and 19 or 18 for the
Cr (III) or Cr (Vl)-amended plants, grown in the field-moist or air-dried soils,
respectively (Table 4.3). This indicates that very little Cr was translocated to the
shoots, although greater translocation with Cr (VI) compared to that with Cr (III)
was observed (Table 4.3). Cr (VI) has probably been reduced in the plant roots
to the less biologically active Cr (III), thereby limiting Cr movement to the shoots.

67

Lytle et al. (1998) also observed Cr (VI) reduction In the fine lateral roots of
Eichhomia crassipes (water hyacinth) after 4 hours of metal exposure. Further,
Shewry and Peterson (1974) hypothesized that Cr is unavailable for transport
probably due to its spatial localization in a specific subcellular compartment in
the root cells (the vacuole), as a tolerance mechanism and a common feature of
metal-stressed plants (McGrath at a!., 2002). Vazquez et al. (1994) observed
preferential accumulation of Zn in the vacuoles of root and leaf epidermal cells of
Thiaspi caerulescens (alpine pennycress). The Ni hyperaccumulator, Thiaspi
goesingense (Austrian mustard), was also found to sequester Ni in the vacuoles
in leaf cells (Kramer et al., 2000).

Plant dry weight was used to calculate an index of Cr tolerance in each
soil and Cr treatment (Table 4.3). For the plants in any of the Cr and soil
treatments, the index of tolerance was greater than 50%, which is generally
considered to be the minimum desired biomass production for the plants
growing in a metal- contaminated site (Chang et al., 1992; Baker et al., 1994b).
Tolerance of Cr was influenced by the form of applied Cr, but not the soil
treatment (Table 4.3). In particular, plants had the index of tolerance of 87 and
88% when grown in the dry and moist soils contaminated with Cr (III), while the
index of tolerance dropped to 68 and 70% in the same soils treated with Cr (VI)
(Table 4.3).

68

The bioaccumulation (or transfer) factors (BCF) were also computed for
roots and shoots in relation to Cr applied to the soils (Table 4.3). In general, the
higher the BCF, the higher the plant uptake of Cr from soil. Plant roots had
higher bioaccumulation factors than shoots. For the roots, the bioaccumulation
factors varied from 2.0 and 2.1 to 3.9 in plants grown in field-moist and air-dried
Cr (III) and Cr (Vl)-treated soils, respectively (Table 4.3). For the shoots, the
highest plant uptake was observed from both field-moist and air-dried Cr (VI)treated soils as compared to those of Cr (III) (Table 4.3). B. Juncea has been
reported to accumulate Cr (Raskin et al., 1994; Salt et al., 1995). However, most
of the studies on Cr accumulation have been performed either hydroponically or
in soils contaminated with high concentrations of anthropogenic or applied Cr.
For example, in an experiment of Kumar et al. (1995), B. juncea had the lowest
bioaccumulation

factor

of

0.1

in

Cr

(III)

treatment

and

the

highest

bioaccumulation factor of 64 in Cr (VI) treatment. Salt et al. (1995) also found
that hydroponically grown B. Juncea has accumulated Cr mainly in the roots with
the ratio of bioaccumulation factor in roots to shoots of 70. Although B. Juncea
plants could accumulate large amounts of Cr in their roots or shoots, they died
within a few days following exposure to 500 mg Cr (VI) kg'”' soil (Shahanden and
Hossner, 2000). Given that Cr added to soil can show several fates including
oxidation, reduction, adsorption, chelation, precipitation as well as remaining in
the solution (Adriano, 1986; Fendorf, 1995), in the present study, solubilization
could be a major problem for B. Juncea to accumulate high Cr concentrations.

69

4.4.2. Chromium influence on plant visible stress

The Cr (VI) treatment resulted in the greatest visible plant stress (Table
4.5). In particular, in both moist and dry soils, some initial stress (between 20
and 50% chlorotic leaves) and moderate stress (between 50% and 80%
chlorotic leaves) were noticed in 78% and 22% of Cr (Vl)-treated plants,
respectively. In contrast, in the Cr (lll)-amended moist and air-dried soils, only
33% and 22% of the plants showed some chlorosis, respectively, while none of
them experienced moderate metal stress. The literature on other plants
illustrates that the effect of Cr (VI) is often more pronounced than that of Cr (III)
(Satayakala and Kaiser, 1993; Jain and Aery, 1998; Mukhopadnyay and Aery,
2000). In particular, chlorosis caused by Cr (VI) was observed by many
researchers (Hara and Sonoda, 1979; Vazquez e ta l., 1987; Davies e ta l., 2001),
which was also found in the present study (Table 4.5). Chlorosis noted in the Cr
(Vl)-treated leaves of B. juncea may be a consequence of toxic effects of Cr (VI)
on the plant roots and stem (Tables 4.8 and 4.10; Figures 4.6, 4.9, and 4.10).
This result is consistent with past studies (Hewitt, 1948; Sharma at a i, 1995;
Jain et a i, 2000). However, probably because only a small amount of Cr
reached the shoots (Tables 4.3 and 4.4), mild chlorosis was observed (Table
4.5).

70

Table 4.5. Visible stress in Brassies juncea after 69 days of growth in Cr (III, VI)-contaminated field-moist and airdried soils.
VISIBLE STRESS (%)
Treatment

mg Crkg'^soii

Dead

Very stressed

Moderate stress

initial stress

Healthy

Control moist

0

0

0

0®

0^

100*

Control dry

0

0

0

0®

0"

100*

Cr (III) moist

100

0

0

0^

33"

67"

Cr (III) dry

100

0

0

0=

22"

78"

Cr (VI) moist

100

0

0

22b

78^

0*

Cr (VI) dry

100

0

0

22b

78^

0*

-

-

-

8.00
(0.015)^

20.34
(0.000)

104.22
(0.000)

F ratio and
(p value)
^ p value.

Within each column, means followed by a common letter are not significantly different from each other using one-way ANOVA
and LSD test (a = 0.05, n = 3).

71

Cr (VI) is also known to alter the content of essential mineral nutrients,
including Fe (Barcelo eta l., 1985), which results in chlorosis. The explanation for
this is that there may be competition between the toxic metal ions and Fe in
enzyme systems involved in chlorophyll formation (Hewitt, 1948). In particular,
the similarity of the ionic radii of Cr (III) and Fe (III) can lead to Cr (III)
substitution for Fe (III) in the proteins, namely catalase and peroxidase, resulting
in loss of their efficiency (Pandey and Sharma, 2002). The first appearance of
chlorosis in the younger leaves of B. juncea may be due to general immobility of
Fe within plants. As a result, the chlorosis mainly affects new growth since even
healthy plants cannot take Fe from older leaves and send it to younger leaves
(Jones, 1998).

4.4.3. Chromium influence on plant root and shoot growth

Plant height was not significantly affected by all Cr and soil treatments
(Figure 4.5). Cell elongation is a complex process involving turgor requirements,
synthesis of wall constituents, and plant growth regulators (Wainwright and
Woolhouse, 1977), namely ethylene, abscisic acid, and gibberellin (Kende e ta l.,
1998). Reduced plant height caused by heavy metals, including Cr, has

72

96
( 0 .8 )

E
o

(0 .2 )

95
( 1 .0 )

( 0 .9 )

( 0 .5 )

94
.5*
5

( 0 .6 )
93

o

o
(A

92
91

Treatment

Figure 4.5. Effect of two chromium species [Cr (III) and Cr (VI)] on shoot height of Brassica juncea grown for 69
days. [Values are means of three replicates. Numbers in parentheses are standard errors. All means are not
significantly different from each other using one-way ANO VA (F ratio = 1.05 and p value = 0.431)].

73

been previously demonstrated, although metal toxicity varied greatly, depending
on soil characteristics and crop type (Kabata-Pendias and Pendias, 1992; Aidid
and Okamoto, 1993; Prasad et al., 1999; Davies et al., 2001). In the present
study, the data on Cr concentration and accumulation in the Cr (III) and Cr (VI)treated plants (Tables 4.3 and 4.4) indicate that low amounts of Cr are
translocated to the shoots, which may be the reason for no significant
retardation of height of B. juncea.

At any time of metal exposure and in both Cr oxidation states, the roots of
B. Juncea were affected to a greater degree than the plant shoots (Figure 4.6).
For example, at 17 days after sowing, root growth in the Cr (lll)-treated moist
and air-dry and Cr (Vl)-amended moist and air-dry plants was reduced by 36%
and 33%, 57% and 54%, respectively, while, in the same Cr and soil treatments,
the shoot growth was decreased by 0% and 1%, 18%, and 17%, respectively, as
compared to the control plants. The plant roots were found to contain much
higher concentrations than the shoots (Table 4.3), which could lead to a
remarkable decrease in their dry weight (Figure 4.6).

Root growth was significantly depressed by both Cr species (Figure 4.6).
The inhibitory effects of Cr on root growth may have resulted from binding of Cr
(III) to the plant tissues and disturbance of osmotic relationships, which lead to
the restricted transport of Ca (II) ions across the plasma membrane into the

74

□ 17 days
@ 36 days
69 days

S 600
£

500

w 200

%

100

/

<<>
c>

Treatment
Figure 4.6. Effect of two chromium species [Cr (III) and Cr (VI)] on mean root
and shoot dry weight of Brassica juncea grown for different exposure periods.
[Within each harvest time, means followed by a common letter are not
significantly different from each other using one-way ANO VA and LSD test (a =
0.05, n = 3)].

75

cytoplasm (Liu et al., 1992). In this way, the level of free Ca (II) ions in the cell
becomes very low, which leads to a failure of calmodulin (CaM ) in activation of a
number of key enzymes, including Ca-ATPase (Liu et a!., 1992). However, at
any exposure period, Cr (VI) appeared to be more toxic than Cr (III). In
particular, at 36 days after planting, for Cr (lll)-treated plants grown in the moist
and air-dry soils, root dry weights were 136 and 135 mg, respectively, whereas,
these values dropped to 83 and 85 mg for Cr (Vl)-treated plants (Figure 4.6).
Since the Cr (lll)-treated plants absorbed a similar amount of Cr as those grown
in the Cr (VI) treatment (Table 4.4), root growth was expected to be similarly
affected by the application of both Cr treatments. However, being a strongly
oxidizing agent, Cr (VI) has been linked to structural and ultrastructural
alterations in plants (Vazquez et a!., 1987). In contrast, Cr (III) is less phytotoxic
due to its lower oxidizing potential (Gauglhofer, 1984).

Over the course of the experiment, the dry weight of the plant shoots
was significantly inhibited by Cr (VI) treatment, while it was not sensitive to Cr
(III) treatment (Figure 4.6). Both higher concentration and accumulation (Tables
4.3 and 4.4), and toxicity of Cr (VI) are likely responsible for this trend.

At any sampling time, there was no significant effect of soil treatment on
root and shoot dry weights (Figure 4.6). This was not expected, particularly at
the first harvest, when oxidation of a small portion of Cr (III) was observed in the

76

field-moist soil (Figure 4.1). Nevertheless, the concentration of formed Cr (VI)
perhaps was not high enough to cause a significant decrease in the dry weight
of roots and shoots.

4.4.4.

Chromium influence on plant root exudation

Capillary electrophoresis (CE) was applied to separate, identify, and
measure low-molecular-weight organic acids in root exudates of B. juncea. This
new technique has been proven to allow for a rapid and efficient separation of
charged compounds present in small sample volumes (Barbas et al., 1998).
Moreover, the separation can often be achieved directly in aqueous media,
without sample pretreatment (Barbas et a!.,

1998).

Electrokinetic sample

injection is known to enhance CE sensitivity (Galli et a!., 2003), which was
crucial for the

low concentrations

of organic acids

in this study.

The

electropherograms of standard mixture of organic acids and solutions of root
exudates, collected from the control and Cr (Vl)-treated plants at three sampling
times, are illustrated in Figures 4.7 and 4.8. It can be seen that malic, citric,
succinic, and acetic acids are consistently present in all samples at any harvest
time. In addition, at any time of Cr exposure, all peaks in the Cr (Vl)-amended
plants are higher than those of the control plants (Figures 4.7 and 4.8).
Moreover, the magnitudes of absorbance seem to decrease from vegetative (17

77

0.04 -

-

0.04

0.03 —

—

0.03

-

0.02

-

0.01

3

(Q
8
C

12

s

S

0.02 -

05

in

€

00

0.00
-

0.04

0.03 -

—

0.03

-

0.02

8C

I
<

I

I

0.00

0.04 -

I 0.02

i

-

o
c

0.01 -

-

0.00

0.01

0.00
7.0

7.1

7.2

7.3

7.4

7.5

7.6

7.7

7.8

7.9

8.0

8.1

8.2

8.3

8.4

8.5

8.6

8.7

8.8

8.9

9.0

9.1

9.2

I

9.3

M ig ra tio n tim e (m in u tes)

Figure 4.7. Electropherograms of: (a) 10 pM standard solution mixture of malic, citric, succinic and acetic acids; (b)
root exudates collected from Brassica juncea at 17 days (electrokinetic injection of 10 kV for 10 sec). Solid and
dashed graphs represent control and Cr (Vl)-treated plants, respectively.
78

3

re
8
C
re

■0s

0 .0 4 -

-

0 .0 4

0 .0 3 —

-

0 .0 3

8

c
0 .0 2

—

-

0 .0 2

0.01

-

-

0.01

<

1

0.00

3

re
8
C

0.00

0 .0 4 -

-

0 .0 4

0 .0 3 —

-

0 .0 3

—

-

0 .0 2

-

-

0.01

I 0 .0 2

I

S

0.01

3
a
8

I

I

0.00

0.00

7 .0

7.1

7 .2

7 .3

7 .4

7 .5

7 .6

7 .7

7 .8

7.9

8 .0

8.1

8 .2

8 .3

8 .4

8 .5

8 .6

8.7

8.8

8 .9

9 .0

9.1

9.2

9 .3

M ig ra tio n tim e (m in u te s )

Figure 4.8. Electropherograms of root exudates collected from Brassica juncea at: (a) 36 days; (b) 69 days
(electrokinetic injection of 10 kV for 10 sec). Solid and dashed graphs represent control and Cr (Vl)-treated plants,
respectively.
79

days) to flowering (36 days) and further to fruiting (69 days) stages of plant
development (Figures 4.7 and 4.8).

Quantitative data, however, were not collected with the electrokinetic
sample injection. Despite the fact that some researchers have determined trace
amounts of anions, including organic acids, in different samples (Ehman et al.,
1997; Dahlen e ta l., 2000; Desauziers et a i, 2000; O ’Flaherty et a i, 2001), this
type of sample introduction often suffers from matrix bias and poor precision,
which was also observed in the present experiment (data not shown), and it is,
therefore, not recommended for quantification (Galli et a i, 2003). Accordingly, a
hydrodynamic or pressure injection, which does not discriminate between the
ions, thereby injecting the same effective sample volume of each ion (Devevre et
a i, 1994), was used for quantification of low-molecular-weight organic acids in
the present study. The reproducibility of the quantitative sample introduction
using this method is shown in Table 4.6. When an intermediate standard (100
pM) was run six times per day, the standard deviation of migration time varied
from 1.2% for citric and acetic acids to 1.3% for malic and succinic acids,
respectively, while the standard deviation values for peak areas were 2% for
citric and succinic acids and 3% for malic and acetic acids, respectively (Table
4.6).

80

Table 4.6. Reproducibility of pressure sample introduction of 100 pmol L'^
standard mixture of malic, citric, succinic, and acetic acids in capillary
electrophoresis.
Organic acid

Citric

Malic

Succinic

Acetic

Migration time (min)
Mean

7.286

7.514

8.009

8.976

STDEV (%)

1.3

1.2

1.2

1.3

Peak area
Mean

6032

7891

6513

5017

STDEV (%)

3

2

2

3

However, the pressure sample injection appeared to be much less
sensitive than the electrokinetic injection (data not shown). Therefore, the
samples were concentrated by freeze-drying, just prior to the runs. The same
trends as in Figures 4.7 and 4.8 appeared in all samples at all harvest times for
three organic acids, namely malic, citric and succinic acids (Figures 4.9 and
4.10). In contrast, acetic acid, detected with the electrokinetic injection (Figures
4.7 and 4.8), seemed to nearly disappear in all samples (Figures 4.9 and 4.10).
Being the most volatile, it had probably been lost during the freeze-drying
procedure. Other sample preparation techniques, for example, use of anion
exchange resins (Robinson, 1986), should therefore be considered for efficient
extraction of volatile organic acids.

81

0 .0 1 2 5

0.0 12 5

0.0100

0.0100

•

0 .0 07 5 -

a
.a

0 .0 05 0

-

0 .0 02 5

.

-

0 .0 0 7 5

.

0.0 05 0

8

S

00

0 .0 0 2 5

0 .0 00 0 -

-

0.0100

I

•

0 .0 0 7 5 -

0 .0 05 0

.

0 .0 02 5

•

0.0 00 0

I

I

0.0 00 0
0 .0 1 2 5

0 .0 12 5

30
1
8
c

8

0.0100

-

0 .0 0 7 5

•

0.0 05 0

8
S

0 .0 0 2 5

<

-

-

7.0

•

7.1

7 .2

7.3

7.4

7.5

7.6

7.7

7.8

7.9

8.0

8.1

8 .2

8.3

8.4

8.5

8.6

8.7

8.8

8.9

9.0

9.1

9.2

0.0 00 0

9.3

M igration tim e (m inutes)

Figure 4.9. Electropherograms of: (a) 100 pM standard solution mixture of malle, cltrlc, succlnlc and acetic acids;
(b) root exudates collected from Brassica Juncea at 17 days (pressure Injection of 0.1 psi for 10 sec). Solid and
dashed graphs represent control and Cr (Vl)-treated plants, respectively.

82

0 .0 1 2 5

0.0 12 5

.

0.0100

3
ra
8
c

I

0.0 0 7 5 -

I

I
<

-

0.0075

0.0050

0.0050

0.0 02 5

0.0025

0.0000

i
8

0.0100

—

-

0.0 12 5

0.0100

0.0100

0.0 07 5 -

0.0075

0.0050

0.0050

0.0025

0.0025

-

—

7.0

7.1

7.2

7.3

7.4

7.5

7.6

7.7

7.8

7.9

8.0

8.1

8.2

8.3

8.4

8.5

8.6

8.7

8.8

8.9

9.0

9.1

9.2

I

I
<

0.0000

0.0 12 5

0.0000

1
8

8

I

0.0000

9.3

M igration tim e (m inutes)

Figure 4.10. Electropherograms of root exudates collected from Brassica Juncea at: (a) 36 days; (b) 69 days
(pressure injection of 0.1 psi for 10 sec). Solid and dashed graphs represent control and Cr (Vl)-treated plants,
respectively.

83

In general, root exudation in the rhizosphere occurs as a result of
mechanical Injury from small Insects and nematodes, growth of lateral roots, and
abrasive action of soil particles (Rovira, 1969). Low-molecular-weight organic
acids released Into soils can also Increase the ability of plants to survive and
grow normally under conditions of nutrient deficiency (Neumann and Romheld,
1999). In particular. Increase In organic acid efflux has been observed under K
(Kraffczyk et al., 1984), Fe (Mulette et al., 1974), and a general nutrient
deficiency (Jones and Darrah, 1995). The organic acid anions In root exudates
can chelate Fe and Mn or lower rhizosphere pH, thus making Mn, Fe, and Zn
more available for plant uptake (Marschner, 1986). Similarly, organic acid anions
can form complexes with Ca and AI present In soil as Insoluble phosphates and
liberate P uptake by roots (Marschner, 1986). In the present Investigation, at any
sampling time or with any Cr and soil treatments, citric acid was the most
abundant of the three organic acids exuded by B. juncea (Table 4.7). The
presence of citric acid In root exudates of other plant species Is considered to be
mostly related to P absorption (Neumann and Romheld, 1999). For example. In
S. napus, a large Increase In succlnlc, malic, and citric acid levels (by 70 and 12
times, respectively, for malic and citric acids) has been observed under P
deficiency (Zhang et al., 1997). However, In the present study, soils were
supplied

with

the

N-P-K

fertilizer.

Therefore,

the

abundant

citrate

84

Table 4.7. Organic acids (|jg L'^) in root exudates of Brassica juncea treated with 100 mg kg'^ of either CrCb 6 H 2 O
or KaCraO?.
Citric

Malic

Organic
acid

F ratio
and
(p value)
time

Treatment

17
days

36
days

69
days

Control
m oist

452*

284"

(3)'

(5)

Control
dry

453*
(3)
455*

285"

131*
(1)
132*

(8)

(5)

293"
(11)

129*
(2)
132*
(2)

389.22
(0.000)

214'

C r (III)
m oist

(4)

298"
(8)
442*

1122.53
(0.000)
520.44
(0.000)

C r (III)
dry

449*
(2)

C r (VI)
m oist

553b

(10)

(4)

(4)

471.44
(0.000)

C r (VI)
dry

549b

(8)

450''
(13)

217'
(6)

215.19
(0.000)

treatment

62.10
(O.OOOf

55.39
(0.000)

21.17
(0.000)

702.56
(0.000)

-

Succinic
F ratio
and
(p value)
time

17
days

36
days

69
days

225.67
(0.000)

474*

315"

144"

(4)

(5)

(6)

370.95
(0.000)

476*
(8)
473*

309"

152"

(7)

(4)

316"
(2)
315"
(3)

146"

17
days

36
days

69
days

714*
(8)
713*
(21)
718*
(13)
711*
(10)

527"
(21)
543"
(15)

224*

539"

230*
(5)
225®
(3)

1271.63
(0.000)
758.08
(0.000)

471*

794b

630''
(19)

339'
(4)
(11)
342'
793f
635''
(18)
(10)
(7)
F ra tio a n d (p value)

215.46
(0.000)

572“’
(6)
571»
(6)

479*

39.56
(0.000)

7.03
(0.003)

(6)

540"
(7)

71.84
(0.000)

(5)

230*
(6)

38.12
(0.000)

219.10
(0.000)

-

(7)
(7)

F ratio
and
(p value)
time
719.67
(0.000)
416.06
(0.000)

(7)

480.16
(0.000)

145"
(8)

447.44
(0.000)
327.62
(0.000)

480*
(5)

268*
(8)
272*
(6)

175.07
(0.000)

64.71
(0.000)

(7)

497.60
(0.000)

-

^ Standard error. ^ p value.
Within each acid, means followed by a common letter are not significantly different from each other using one-way ANOVA and
LSD test (a = 0.05, n = 3).
85

production could not be related to the low P concentration in the studied soils.
On the other hand, the amount of other nutrients, Fe, for example, which
appears to be limited in the soils under present investigation (Pep = 0.68%:
Arocena and Sanborn, 1999), might have been insufficient for normal growth of
B. juncea, thereby enhancing citric acid exudation. There are reports in the
literature concerning changes in concentrations of organic acids in the roots of
several plant species induced by Fe deficiency. As an example, citrate and to a
lesser extent malate increased with Fe deficiency in roots of H. annuus (Venkat
Raju et al., 1972). Citric and malic acids are not the only organic acids which
exudation is enhanced in Fe-deficient roots. For instance, Alhendawi et al.
(1997) found that increase in bicarbonate in nutrient solution led to chlorosis and
to increase of citrate, malate, aconitate, and succinate in the roots of H. vulgare.
Sorghum b/co/or (sorghum), and Z. mays.

The solubilizing ability of organic acids has been reported to be parallel to
their metal-binding capacity, which, in turn, is correlated with their dissociation
constants (Srivastava et al., 1998a). The order of the excretion of the organic
acids obtained in the present study (Table 4.7) is in agreement with the
dissociation constants, i.e., Ka^, Ka2 , for citric acid (7.45x10’^, 1.73x10'®), malic
acid (3.48x10'"^, 8.0x10'®), and succinic acid (6.21x10'®, 2.31x10'®) (Martell and
Smith, 1976).

86

Table 4.7 also demonstrates that, at any collection time, exposure of
plants to Cr (VI) caused a significant increase in concentration of each organic
acid, while Cr (III) treatment did not affect root exudation of B. juncea. Organic
acids such as citric, malic, and aspartic acids in root exudates of Z

mays

(Srivastava et al., 1998a), citric, oxalic, and aspartic acids in Lycopersicon
esculentum (tomato) (Srivastava at a!., 1998b), and oxalic, malic, and glycine in
Triticum vulgare (brad wheat) (Srivastava at a!., 1999), have been reported to
have a potential for reducing Cr (VI) and chelating either formed or present Cr
(III). As a result, enhanced root exudation, observed in this study with Cr (VI)
treatment, could be a defense (detoxification) mechanism of the plant. On the
other hand, the concentrations of organic acids (Table 4.7) seem insufficient for
the binding of each of the Cr (III) and Cr (VI) present in the roots (Table 4.3).
This could be due to utilization of some portion of root exudates by rhizospheric
microorganisms, which may have still been adhered to the roots after a gentle
wash and sonication. As Cr (III) did not increase exudation of organic acids,
while Cr (VI) did, it is also possible that there was simple leakage of organic
acids through ruptured plasma membranes, caused by high oxidative potential
of Cr (VI).

In all treatments, the highest levels of any acid were detected at the
vegetative stage (17 days after planting). These levels decreased towards
flowering and fruiting stages (36 and 69 days after planting, respectively) of plant

87

development (Table 4.7). Seedling development has a high nutrient demand not
only for growth of Individual organs, synthesis of new cytoplasm and sub-cellular
organelles and cell walls, but also for cell division and expansion (Moorby and
Besford, 1983). In contrast, during the reproductive stage, growth of plant slows.
Furthermore, at maturity, the plant does not produce much chemical energy
(photosynthetic rates slow down) and requires very little nutrients (Spaugh,
1999). This is one possible explanation why root exudation declined with time
(Table 4.7). These findings are in agreement with a previous study of Lucas
Garcia et al. (2001), who observed 50 and 80% decrease in the total amount of
organic acids exuded by Lupinus albus (white lupine) and Lupinus iuteus
(European yellow lupine), respectively, at the fruiting stage compared to that at
the flowering stage of plant development. Further, the rate of leakage of oxalic
and succinic acids was greater from the youngest seedlings than from older
mycorrhizal and non-mycorrhizal plant species (Schwab eta!., 1983).

4.5.

Microscopic effects of chromium on plant growth

4.5.1. Chromium influence on shoot anatomical characteristics

The stem diameter significantly decreased in Cr (VI) treatment with a
reduction in the width of primary xylem and number of both vascular bundles
and xylem cells (Table 4.8). A small amount of Cr, either soluble Cr (VI) or

88

Table 4.8. Effect of two chromium species [Cr (III) and Cr (VI)] on number of vascular bundles and xylem cells,
stem diameter, and width of epidermis, cortex, primary (1°) and secondary (2°) phloem and xylem, and pith in
Brassies juncea after 69 days of growth.
Width (%)

Num ber
Stem
diameter
(mm)

vascuiar
bundles

xylem
cells

Epidermis

C ontrol m oist

2.6»
(0.0)1

37»
(1.2)

14»
(0.0)

1.5»
(0.0)

C ontrol dry

2.7»
(0.0)

36»
(0.7)

13»
(0.8)

C r (III) m oist

2.6»
(0.0)

35»
(0.7)

C r (III) dry

2.6»
(0.0)

C r (VI) m oist

C r (VI) dry

Treatment

Xylem

Pith

1°

2°

1°

6.6»
(0.1)

2.8»
(0.2)

1.8»
(0.2)

15»
(0.2)

9.0»
(0.3)

64»
(0.6)

1.7»
(0.2)

6.2»
(0.4)

2.8»
(0.2)

1.8»
(0.2)

14»
(0.1)

9.0»
(0.4)

64»
(0.2)

13»
(0.7)

1.4»
(0.2)

6.9»
(0.5)

2.7»
(0.2)

1.5»
(0.0)

15»
(0.2)

8.5»
(0.3)

64»
(1.0)

35»
(0.5)

15»
(0.7)

1.5»
(0.0)

7.2»
(0.5)

2.8»
(0.2)

1.8»
(0.2)

14»
(0.1)

9.2»
(0.3)

63»
(0.5)

2.2^
(0.0)

19"
(0.5)

11"
(0.3)

1.8»
(0.0)

8.0»
(0.3)

3.4»
(0.2)

2.2»
(0.5)

9.2"
(0.4)

10»
(0.1)

65»
(0.5)

2 Jb

16"
(1.4)

10"
(0.5)

1.9»
(0.0)

7.3»
(0.9)

3.2»
(0.2)

1.9»
(0.0)

9.2"
(0.9)

11»
(0.7)

66»
(0.8)

67.57
(0.000)

7.05
(0.003)

1.87
(0.174)

0.73
(0.612)

0.37
(0.859)

0.30
(0.905)

41.42
(0.000)

1.53
(0.252)

1.75
(0.198)

(0.0)

F ratio and
(p value)

Phloem

Cortex

43.29
(0.000/

2°

^ S tandard error. ^ p value. W ithin each colum n, m eans follow ed by a com m on letter are not significantly different from each
o ther using one-way AN O V A and LSD te st (a = 0.05, n = 3).

89

reduced Cr (III), can be transported to the upper plant parts by an active
mechanism via the transpiration stream in xylem or by a mechanism similar to
that of Ca^"' (Skeffington et al., 1976; Barcelo et al., 1985). Toxic concentrations
of Cr decrease the number and diameter of xylem cells (Barcelo and
Poschenrieder, 1990). In a similar study, the sizes of both phloem and xylem
cells decreased in stems of Cr (Vl)-treated P. vulgaris (Vazquez et al., 1987).
Suseela et al. (2002) also observed a decrease in the number of fibers in the
shoot of S. lacustris, treated with 8 mg L'^ Cr (VI) for 30 days, as an indicator of
pollution with Cr (VI). The inhibition of these plant characteristics may be a result
of interference of Cr (VI) with cell division and cell elongation that leads to
decrease in cell water content (Barcelo and Poschenrieder, 1990). Heavy metals
have been shown to affect both processes in plants. This could be due to
insufficient supply of nutrients (Barcelo et al., 1985) and plant growth regulators
from the affected roots, which may in turn influence the differentiation of tissues
in stems (Setia and Bala, 1994). For example, limited supply of cytokinins, which
are mainly synthesized in roots (Van Staden and Davey, 1979), has been linked
to inhibition of lateral shoot development during AI toxicity stress (Pan et al.,
1988).

The leaves were not affected by any Cr and soil treatment (Table 4.9).
Some changes in the plant leaves were expected, since some visual stress was

90

Table 4.9. Effect of two chromium species [Cr (III) and Cr (VI)] on leaf thickness, thickness of palisade and spongy
mesophyll, palisade cell layer number, and leaf vascular bundle number of Brassioa juncea after 69 days of growth.

Leaf thickness
(pm)

Palisade mesophyll
thickness
(pm)

Spongy mesophyll
thickness
(pm)

Num ber of
palisade cell
layers

Number of
vascular
bundles

Control moist

379
(4)1

160
(9)

167
(8)

4
(0.0)

2
(0.0)

Control dry

377
(0.0)

161
(2)

167
(5)

4
(0.2)

1
(0.2)

Cr (III) moist

376
(6)

168
(7)

161
(5)

4
(0.2)

2
(0.5)

Cr (III) dry

361
(10)

166
(7)

168
(3)

4
(0.0)

2
(0.0)

Cr (VI) moist

369
(6)

157
(5)

167
(2)

4
(0.0)

1
(0.5)

Cr (VI) dry

366
(13)

154
(11)

161
(4)

4
(0.0)

2
(0.2)

F ratio and
(p value)

0.43 (0.817/

0.26 (0.928)

0.23(0.944)

1.20(0.366)

0.88(0.523)

Treatment

^ Standard error,

p value.

Within each column, means are not significantly different from each other using one-way AN OVA (a = 0.05, n = 3).

91

observed in the Cr-treated plants (Table 4.5). However, only one leaf, at the 3'"'^
node, was studied. This suggests that Cr-injured or relatively immature leaves
should also be collected, since an individual leaf may not be representative of
leaves of the entire plant.

4.5.2.

Chromium influence on root anatomical characteristics

The results indicate that tap and lateral roots of 6. juncea were sensitive
to Cr, although soil treatment did not significantly affect any anatomical
characteristics of the roots (Table 4.10). Roots are generally the first organs to
contact toxic metals in soils and they usually accumulate significantly higher
amounts than the aerial plant parts (stems and leaves) (Breckle, 1989). In the
present study, most of the Cr was taken up and retained by the plant roots
(Tables 4.3 and 4.4), which may also explain why the cells of the plant leaves
did not show any metal injury (Table 4.9). In past reports on Cr in plants, it has
been hypothesized that the reduction of toxic Cr (VI) to less toxic Cr (III) occurs
in the roots (Parr and Taylor, 1980; Micera and Dessi, 1988; Lytle at a/., 1998;
Aldrich at a i, 2003). However, when the Cr (VI) concentration within cells
exceeds the reducing capacity of cells, toxicity might occur (Vazquez at a i,
1987; Kortenkamp at a i, 1991). On the other hand, the amount of Cr (VI) formed
in the Cr (lll)-treated field-moist bulk soil (Figure 4.1) was probably insufficient to

92

Table 4.10. Effect of two chromium species [Cr (III) and Cr (VI)] on root diameter, xylem diameter, number and
width of all cells In a part of xylem and large cells In entire xylem of tap and lateral roots of Brassies juncea after 69
days of growth.

Tap

Lateral

Tap

Lateral

Lateral

Tap

Lateral

Tap

Lateral

root

root

root

root

root

root

root

root

root

root

root

root

187®

102®
(2)
112®
(20)
112®
(11)
110®
(0.0)
73"
(0.0)
73"

20®
(0.0)
22®
(3)
19®

43®

16®
(0.3)
16®
(0.0)
20®

17®
(0.0)
18®

12®
(2)
12®

(2)
12"

(1)
10®

(1)
20®

(1)
19®

(1)
13"

(1)
9®

(1)
20®

(1)
9®

(1)
8®

(1)
20®

(1)
9®

(1)
7®

(1)

(0.0)

4®
(0.3)
3®
(0.0)
2®
(0.3)
3®
(0.0)
2®
(0.4)
2®
(0.5)

36®

(3 ),
132^
(8)

909®
(16)
883®
(10)
644b
(80)
616^
(57)
396®
(25)
465®
(16)

6®
(0.5)
6®
(0.3)

C r (VI)
m oist
C r (VI)
dry

1044®
(29)1
1017®
(16)
744b
(24)
766f
(20)
622®
(33)
706®
(14)

F ratio and
(p value)

35.34
(0.000)^

15.63
(0.000)

16.32
(0.000)

3.43
(0.046)

8.39
(0.001)

1.75
(0.211)

Control
m oist
C ontrol
dry
C r (III)
m oist
C r (III) dry

(1)
192®
(0.4)
194®
(5)
185®
(9)
131^

Number of large
cells in entire
xylem

Width of cells in
part of xylem
(pm)

Tap

Treatment

Xylem
diameter
(pm)

Number of cells
in part of xylem

Width of large
cells in entire
xylem
(pm)
Tap
Lateral

Root
diameter
(pm)

4^
(0.0)
4"
(1.0)
2b
(0.5)
2b

(1)
41®
(2)
32'*
(0.0)
33 "

(1)
45®
(1)
63"

(1)

(1)

(1)
61"
(3)
97®
(3)
92®
(7)

19.00
(0.000)

1.53
(0.265)

28.22
(0.000)

(1)
28®

(1)
16®

(1)
29®

(1)
17®

(1)

(1)

(1)

2.80
(0.078)

9.08
(0.001)

1.49
(0.278)

^ S tandard error. ^ p value. W ithin each colum n, m eans follow ed by a com m on letter are not significantly different from each
oth e r using one-w ay A N O V A and LSD test (a = 0.05, n = 3).

93

significantly inhibit any anatomical characteristics of either tap or lateral roots
(Table 4.10).

The tap or primary roots seemed to exhibit changes in both Cr (III) and Cr
(VI) treatments with the latter being more toxic to the plants than the former
(Table 4.10, Figure 4.11). In particular, the root and xylem diameters were
considerably reduced in the plants grown in the Cr (VI) treatment compared to
those in the Cr (III) treatment; the control plants had significantly lower values
compared to those treated with Cr (III). Moreover, the number of large cells in
the entire xylem was significantly higher for the control plants compared to the
Cr (III, Vl)-treated plants, while the width of these cells was significantly lower
for the Cr (VI) plants compared to both control plants and those grown in Cr (III)
treatment. The similar trend was observed for the width of cells in a part of
xylem; however, the number of these cells was significantly higher in the Cr (VI)treated plants compared with either the control or Cr (lll)-treated plants (Table
4.10). The higher toxicity effect of Cr (VI) on plant roots was expected. It is
known that B. juncea can absorb both Cr (III) and Cr (VI); however, the latter is
taken up more easily and in higher concentrations than the former (Kumar et al.,
1995; Shahandeh and Hossner, 2000). In the present study, the root growth was
severely inhibited in the Cr (Vl)-treated plants (Figure 4.6), thus possibly
lowering the capacity of the plant to take up water from the soil. In general, the
primary toxic effects of heavy metals are their influence on membrane function.

94

200u«

Figure 4.11. Effect of Cr on tap root anatomy. Longitudinal sections (100x) of tap roots from; (a) control, (b) Cr (III)treated, and (c) Cr (Vl)-treated Brassica juncea after 69 days of growth in field-moist soils. Note the smaller root
and xylem diameters and a number of large xylem cells in (b) and further reduction of these root characteristics in
(c) in comparison to (a). Cr (VI) treatment (c) also caused the greatest increase in the number of cells in the part of
xylem (chosen for measurements) compared to either the control (a) or Cr (III) treatment (b).
95

ion balance and enzyme activity, whicti could bring substantial alterations of
water relations at

both the cellular and the whole plant level (Barcelo and

Poschenrieder, 1990). Cell wall extensibility and thus cell expansion have been
reported to be severely inhibited due to metal-induced decrease of cell wall
synthesis (Barcelo and Poschenrieder, 1990). For example, Cr (VI) has been
found to alter Golgi activity (Vazquez et al., 1987).

Being a strongly oxidizing

agent, Cr (VI) appears to damage various cellular components including cell wall
and membranes resulting in their structural change (Vazquez et al., 1987). This
may in turn reduce membrane water permeability and, as a result, contribute to
reduced water uptake. Decrease of vessel diameter in plants is also one of the
effects caused by Cr toxicity (Barcelo and Poschenrieder, 1990). There are
several reports on the influence of Cr on the vascular system of roots. For
example, the vessel density, the dimension of vessel elements and number of
fibers all have decreased significantly in the root of S. lacustris treated with Cr
(VI) (Suseela et al., 2002). In the present study, the narrower xylem elements in
the Cr (III, Vl)-treated plants compared to elements in the control plants may
have lead to decrease in root and xylem diameters. It is generally suggested that
the growth of the stelar or vascular tissue of the roots is regulated by the activity
of meristematic tissue and plant growth regulators (Burstrom and Svensson,
1972). Cytokinins and auxins are known to promote xylem differentiation
(Dalessandro and Roberts, 1971; Dalessandro, 1973; Minocha and Halperin,
1974; Aloni, 1987). Aloni (1987) proposed a hypothesis according to which the

96

rate of conduit, i.e., production of xylem vessels, is positively correlated with the
amount of auxins that the differentiating cells receive. Moreover, the final size of
a conduit is determined by the rate of cell differentiation. Cell expansion ceases
after the secondary wall is deposited; therefore, rapid differentiation results in
narrow vascular elements, while slow differentiation permits more cell expansion
and therefore results in wide vascular elements. Conduit density is also
controlled and positively correlated with auxin concentration (Pizzolato, 1982;
Aloni and Zimmermann, 1983). Auxins probably do not act alone, but interact
with cytokinins, which generally stimulate cell division (Boote, 1977). The results
in this study suggest that Cr (VI) decreased the size, but increased the density of
xylem cells perhaps through high concentrations of auxins and cytokinins. In
addition, the high density of the very narrow xylem cells in the Cr (III, Vl)-treated
plants in this study may be a defensive response of B. juncea to metal stress.
However, there is a lack of experimental data on this topic and further studies
are required to investigate this phenomenon.

In the lateral roots, the values for root and xylem diameters were
significantly lower for the Cr (Vl)-treated plants, while remained similar for those
exposed to Cr (III) compared to control (Table 4.10 and Figure 4.12). However,
the number and the width of the large cells in the entire xylem, the number of
cells in a part of xylem, and the width of these cells were not significantly

97

^

TOO u m

^

100 um

TOO u m

Figure 4.12. Effect of Cr on lateral root anatomy. Longitudinal sections (200x) of lateral roots from: (a) control, (b)
Cr (lll)-treated, and (c) Cr (Vl)-treated Brassica Juncea after 69 days of growth in field-moist soils. Note the
decrease in root and xylem diameters in Cr (VI) treatment (c) in comparison to control (a) and Cr (III) treatment (b)
and the absence of any significant effect of both Cr treatments on the number and width of large xylem cells in the
entire xylem and in the part of xylem chosen for measurements.
98

affected by any Cr treatment (Table 4.10). A major function of lateralroots

is

believed to be nutrient and water uptake as well as mycorrhizalformation, while
a major function of tap roots is primarily mechanical support and transport of
water to shoots (Raven et al., 1999). Therefore, higher Cr toxicity would be
expected in the lateral rather than in the tap roots. However, several reports in
the literature have demonstrated the extremely high resistance of lateral roots to
heavy metals (Ivanov, 1994; Seregin and Ivanov, 1998). Researchers believe
that this phenomenon is due to the barrier properties of the endodermis in lateral
roots compared with that in tap roots. In particular, while the transport of heavy
metals across the endodermal barrier, i.e., the Casparian strip, is hampered.
Moon (1986) found that laterals can interrupt the continuity of the endodermis for
a brief time, thereby forming gaps for metal transport (Wierzbicka, 1987; Ksiazek
and Wozny, 1990; Seregin and Ivanov, 1997). Therefore, due to more limiting
transport of Cr to the root vascular system and further to shoots, in this
experiment, the tap root cells might have had higher Cr concentrations than the
lateral root cells. As a result, the former would be more seriously damaged than
the latter, which was indeed the case (Table 4.10).

4.5.3. Chromium spéciation and distribution within plant

Figure 4.13 shows the XANES data and LC-XANES fittings for the Cr
(Vl)-treated root and leaf of B. juncea grown in field-moist soil for 69 days. The
lack

of

a

pronounced

pre-edge

peak

of

Cr

(VI),

present

in

the

99

5

3

é

c
0

■l

1

C r ( V l ) - t r e a te d l e a f
1-

LC-XANES

<

■o

fitting

I

«s
Ë

i

0.

C r ( V l ) - t r e a te d r o o t
LC-XANES

fitting

Photon energy (keV)

Figure 4.13. XANES spectra and LC-XANES fittings of Cr (Vl)-treated leaf and root of Brassica juncea grown for 69
days in the field-moist soil.

100

reference compound of Cr (Vl)-dichromate (Figure 4.2), Indicates that, in both
the root and leaf, the Cr exists as Cr (III). Therefore, B. juncea converted the
more toxic Cr (VI) into the less toxic Cr (III). The conversion clearly occurred in
root tissues, since most of Cr (III) was observed in the plant roots (Figure 4.13).
In studies that used similar XAS technique, both the lateral roots and leaves of
E. crassipes, supplied with Cr (VI) in nutrient solution, contained only Cr (III)
(Lytle at a/., 1998). In addition, Zayed at a i (1998) reported the absence of Cr
(VI) species in the roots of 4- to 5-week-old B. olaracaa and Brassica rapa
(turnip). Yet in another study, the XAS results demonstrated that Cr (VI), taken
up by the roots of Prosopis sp. (mesquites), was fully reduced to Cr (III) and
transported to the leaf of the plant (Aldrich at a!., 2003).

Skeffington at a i (1976) studied Cr uptake and transport in H. vulgara
seedlings. The authors hypothesized that Cr is transported largely via xylem.
The more mobile Cr (VI) apparently moves more easily than the less mobile Cr
(III) due to retention of the latter by ion exchange on vessel walls (Skeffington at
a i, 1976), as it happens for Ca (II) (Shewry and Peterson, 1974). Cr (lll)-organic
acid complexes are water-soluble and, thus, are mobile. They are not retarded
by ion exchange and, therefore, move more quickly than Cr (III) in roots and
shoots. The mechanism(s) by which plants can reduce Cr (VI) to Cr (III) may be
related to the formation of such complexes. For example, Aldrich at a i (2003)
found high amounts of Cr (III) acetate (about 58% ) in the roots of Prosopis spp.

101

Due to root respiration, the amount of CO 2 at the root surface is generally high
(Paul and Clark, 1989). The authors therefore hypothesized that Cr (VI) was
possibly reduced via an oxidation-reduction reaction with an acetate-type ligand,
given the high negative potential of the redox pair of COz/acetate (Aldrich et al.,
2003). In another study (Lytle at a!., 1998), it was also found that Cr was bound
to oxalate, a low-molecular-weight ligand. B. juncea might have the similar
mechanism of Cr (VI) reduction via ligand formation with a low-molecular-weight
organic acid. The XANES data on the fitting of Cr (III, VI) model compounds are
illustrated in Table 4.11. In the root samples of the Cr (Vl)-treated plants, from
the reference compounds used in the present study, the best fits were found
with Cr (III) as Cr (lll)-acetate present at 72% as well as with a mixture of Cr (III)oxalate (14%) and Cr (lll)-formate (9%). A small amount of Cr (VI) as KaCraOr
(5%) was also present in the roots (Table 4.11), thereby confirming that nearly
complete conversion of Cr (VI) to Cr (III) occurred. One possibility of how Cr (III)
enters the roots is that secreted acetic acid, which was qualitatively identified in
root exudates of B. Juncea (Figures 4.7 and 4.8), could have reduced Cr (VI)
externally and then chelated Cr (III) may have been taken up by the roots, while
some Cr (VI) could have been transported directly into the root cells. However,
quantitative data on acetic acid is needed to confirm this. Additionally, the
presence of low concentrations of other organic acids detected in the root
exudates (Table 4.7) suggests that their accumulation could not confirm the
observed Cr (VI) reduction. In this study, roots were not analyzed for internal

102

organic acids that are found to be quite different in quality and quantity from the
root exudates (Gleba et al., 1999). Therefore, the concentrations of the internal
organic acids might have been sufficient enough to promote the Cr (VI)
reduction.
Table 4.11. Distribution of chromium compounds (%) in a root and leaf of
Brassica juncea grown for 69 days in field-moist soil treated with 100 mg kg'”'
of K2 Crz0 7 (n = 2).
Reference compound

Root

Leaf

Cr (Vl)-dichromate

5

0

Cr (lll)-chloride hexahydtared

0

0

Cr (lll)-formate

9

0

Cr (lll)-acetate

72

19

Cr (lll)-trioxalate

14

81

In the leaf tissues, Cr (lll)-oxalate (81%) and Cr (lll)-acetate (19%)
species were detected (Table 4.11). These findings suggest that Cr (III),
chelated by acetate externally in the plant rhizosphere, could have been
transported to the leaf cells to form a complex with internal oxalic acid in the
vacuoles, which are well-known storage compartments of large amounts of
organic acids in plants (Hodgkinson, 1977). On the other hand, if Cr (III) was
chelated by acetate internally, it could have moved from the cytosol of the root
cells, which occupies a small proportion of the cell volume (Rauser, 1999), into
the major part of root cells, i.e., the vacuole, to produce Cr (lll)-oxalate and
release acetate for return to the cytosol. Nevertheless, future studies on internal

103

concentrations of organic acids, including oxalic and acetic acids, in root and
leaf tissues as well as on distribution of these organic acids between the cytosol
and the vacuoles are required to confirm these speculations.

The distribution of Cr (white color) in a cross-section of a leaf and a
longitudinal section of a lateral root is presented in Figures 4.14 and 4.15. The
X-ray image of the Cr (lll)-treated leaf shows enriched Cr accumulation in the
lower spongy mesophyll and the abaxial epidermal cells as opposed to that in
the

adaxial

epidermal

and

the

palisade

mesophyll

cells

(Figure

4.14).

Preferential localization of Cr inside the hairy parts of the leaves (trichomes) has
been observed in L thdentata (Arteaga et al., 2000). Corradi et al. (1993) also
found that in the seedlings of S. sclarea, Cr accumulated in relatively large
amounts in epidermal hairs of the cotyledons. Many hyperaccumulator plants
are reported to sequester toxic metals in the vacuole of their epidermal cells
(McGrath et al., 2002). The ability of the leaves of B. juncea to accumulate Cr
preferentially in the epidermis and

lower spongy mesophyll

might have

increased the plant’s tolerance to the metal by protecting photosynthesis, which
takes place predominantly in palisade cells.

The X-ray image of a lateral root tissue (Figure 4.15) illustrates that Cr is
mostly concentrated in the epidermis and cortex compared with the root vascular

104

Chromiiim

I

1CXJ u rri

Figure 4.14. X-ray microprobe image (200x) of a cross-sectioned leaf of Brassica juncea grown for 69 days in fieldmoist soil treated with 100 mg kg CrCl3 6H 20 (Uep = upper or adaxial epidermis, Lep = lower or abaxial
epidermis, Pm = palisade mesophyll, Sm = spongy mesophyll). Note the accumulation of Cr (white color) in the
lower spongy mesophyll and epidermis.

105

Figure 4.15. X-ray microprobe image (200x) of a longitudinal-sectioned lateral root of Brassica juncea grown for 69
days in field-moist soil treated with 100 mg kg'”' CrCl3 6H20 (£ c = epidermis and cortex, Vs = vascular system:
xylem and phloem). Note the greater accumulation of Cr (white color) in the interior epidermis and cortex compared
with the xylem and phloem.

106

system (phloem and xylem). In general, due to the barrier in the endodermis,
i.e., the Casparian strip, epidermal and cortical cells contain higher amount of
metals than the cells in the vascular tissue (Barcelo and Poschenrieder, 1990).
For example, Seregin and Ivanov (1997) showed that in longitudinally dissected
roots, both Pb and Cd entered only one or two layers of the vascular system,
while in the cortex, these ions moved across several layers as far as the
endodermis. The Casparian strip is a lignified part of the primary cell wall
consisting of cellulose and lignin (Schreiber et al., 1994). The plasmalemmae of
endodermal cells are very tightly bound to the Casparian strips so that even
severe plasmolysis does not separate the endodermal protoplasts from the
anticlinal cell walls in this area (Bonnett, 1968). Therefore, the movement of
metal ions from the cortex into the central part of the root can only be
symplastically across the plasma membranes of the cortical and endodermal
cells. The frequent occurrence of plasmodesmata in the periclinal cell walls of
the endodermis should also significantly facilitate the uptake of ions from the
cortical symplast into the endodermal symplast (Clarkson, 1991).

Vazquez et al. (1987) observed both the injury (severely damaged
epidermal cells) and the higher Cr content on the root surface of Cr (Vl)-treated
P. vulgaris compared with the plant cells in the central part of the root, which is
in agreement with the present results. Further, Shewry and Peterson (1974)
found that precipitation of Cr in vacuoles of root cortical cells, which generally
contain relatively large vacuoles (Esau, 1977), may be responsible for the lower

107

injury observed in the central part of the root compared with the epidermal and
cortical cells. Some Cr found in the vascular tissue of the lateral root of B. juncea
(Figure 4.15) could have moved either symplastically to avoid the Casparian
strip or apoplastically through the gaps in the endodermis of lateral roots.

108

5.0.

CONCLUSIONS

The findings in the present study clearly illustrate the effects of the two
forms of Cr on growth of Brassica juncea and soil microorganisms. In particular,
both Cr (III) and Cr (VI) reduced plant root biomass and had a negative effect on
root anatomical properties such as root and xylem diameters and number of
large xylem cells and their width, although this effect was more pronounced in
the tap than in the lateral roots. However, Cr (VI) seemed to affect these plant
characteristics to a greater extent than Cr (III). Despite the fact that leaf cells
were not affected by Cr (VI), this form of chromium was more toxic, causing mild
leaf chlorosis, evident reduction in shoot biomass, increase in root exudation of
low-molecular-weight organic acids, and damage of soil microorganisms,
whereas Cr (III) did not appear to affect any of these plant and soil properties.
Moreover, although neither Cr (III) nor Cr (VI) did inhibit shoot length, the latter
treatment did have a negative impact on the girth of the plant stem and on the
development of the vascular system, particularly the number of vascular bundles
and xylem cells.

The differential response of microorganisms to Cr (III) and Cr (VI) could
be related to contrasting bioavailability of the two Cr forms in the soils. Due to
significantly higher amounts of soluble and, hence, bioavailable Cr (III) and Cr
(VI) extracted from Cr (Vl)-treated soils, soil microbial activity (C flush) was
inhibited in this treatment. The differences in growth responses of 6. juncea to

109

Cr (111) and Cr (VI) suggest that the metal absorption and transport within the
plant may be considerably different for these two Cr species. In particular,
consistent with the literature, B. juncea concentrated Cr mainly in the roots,
while Cr translocation to the shoots was generally low. Moreover, although in the
roots, a similar amount of Cr was accumulated from both Cr (III) and Cr (VI)
treatments, more Cr was translocated to the plant shoots in the latter treatment
than in the former.

Although there were observed injuries in the roots and shoots, B. Juncea
was tolerant of Cr at the concentrations used in the present study. Furthermore,
the XANES analyses detected Cr present mostly as Cr (lll)-acetate and Cr (III)oxalate, in the roots and leaves of the Cr (Vl)-treated plants, respectively. This
demonstrates the ability of S. juncea to convert more toxic Cr (VI) to less toxic
Cr (III) and indicates the important role of organic ligands, most likely originating
from plant root and/or microbial exudates, in plant survival under Cr stress.
Additionally, the several sites of Cr localization at the cellular levels were
identified as metal deposition, and probably sequestration, in epidermal and
cortical cells in the roots and epidermal and spongy mesophyll cells in the
leaves, most likely in the vacuoles.

In summary, despite the fact that, under greenhouse conditions, B. juncea
did not seem to remove much Cr from the soils (0.3 and 0.4% from the Cr (III)

110

and Cr (VI) treatments, respectively), the ability of the plant to tolerate and
reduce

Cr

(VI)

to

Cr

(III)

could

make

it

a

potential

candidate

for

phytostabilization.

Ill

6.0.

FUTURE STU D IES AND RECOM M ENDATIONS

Considering limited Cr bioavailability and low plant uptake, it appears that
the type of soil used in this experiment does not pose significant risk to the
environment. Nevertheless, to estimate biovailability and consequently resolve
the fate and threat of soil Cr, the mass balance studies accounting for leached,
exchangeable,

organically-bound,

Fe/Mn

oxide-bound

and

residual

concentrations of the added metal are needed. Moreover, the conditions of the
Cr-spiked soils used in this greenhouse study might not reflect real conditions in
field soils, either naturally or anthropogenically contaminated (aged) with Cr,
where complex interactions with several other metals are likely. Therefore, field
work is required to receive an accurate assessment of the potential of Brassica
Juncea for remediation of soil Cr. Future research should focus on soil
characteristics such as pH, organic matter, moisture, and mineralogy (clay
minerals, Fe and AI oxides), which are crucial in Cr spéciation and behavior in
soil and, as a result, in plants. The investigation of the effects of higher Cr
concentrations, which could possibly induce metal uptake by B. juncea, should
also be carried out to further elucidate the potential of the plant to tolerate and
remediate soil Cr. Finally, more samples of root, leaf, and stem tissues should
be collected in order to obtain more representative data on Cr distribution in the
plant.

112

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APPENDIX
Table 1. Profile description of soil from Aleza Lake (Elevation 704 m; North
54 03.844’; W est 122 02.473’); Glaciolacustrine parent material.
Horizon

Depth (cm)

Description

L

3.5-1.5

Needles, deciduous leaves, and twigs.

F

1.5-0

Dry; very dark grayish brown (1OYR 3/2);
structureless; loose; fibrous; abundant, fine
to medium roots; common fauna; few
earthworms, larvae; common, random
mycelia; abrupt, smooth boundary;
0.5-2.5 cm thick.

Ahe

0-5.5

Grey (7.5YR 5/1 d); sand; weak, fine,
granular; loose; plentiful, fine to medium
roots; clear, smooth boundary; 1.5-7.0 cm
thick.

Bf

5.5-30.5

Dark yellowish brown (10YR 3/6 m);
sand; structureless; loose; plentiful, fine
to medium roots; clear, smooth boundary;
25.5-28.5 cm thick.

Bfj

30.5-44

Dark yellowish brown (10YR 3/4 m); sand;
structureless; very friable; plentiful, very
fine roots; clear, smooth boundary;
19.0-43.5 cm thick.

BC

44-59.5

Dark brown (1 OYR 3/3 m); sand;
structureless; very friable; few, medium roots;
clear, smooth boundary; 16.5-28.0 cm thick.

C

59.5-81

Very dark grayish brown (10YR 3/2 m);
sand; structureless; very friable; few,
medium roots, abrupt, irregular boundary;
5.5-23.0 cm thick.

lie

81+

Brown (10YR 4/2 m); silty clay; massive;
firm; no roots; common, thin to moderately
thick, on ped surfaces (7.5YR 4/4 m) clay
films.

132