Class: Dicotyledonae
Sub Class: Polypetalae
Series: Thalamiflorae
Order: Parietales
This family is commonly called as 'mustard family'. It includes about 300 genus and about 3700 species. The members have a cosmopolitan distribution. There are around 150 species in India.
Characteristic Features
Habit: Usually annual or perennial herbs.
Root: Taproot system.
Stem: Herbaceous, erect, branched.
Leaves: Simple, alternate, radical or cauline, usually entire, sometimes lobed, petiolate, exstipulate reticulate venation.
Inflorescence: Raceme or corymbose raceme.
Flower: Ebracteate, pedicellate, mostly actinomorphic, bisexual, heterochlamydeous, dimerous or tetramerous hypogynous.
Calyx: Sepals 4, polysepalous, in two whorls of two each imbricate aestivation.
Corolla: Petals 4, arranged in single whorl alternating with sepals, polypetalous, often with long claws and spread out to form a cross. Hence, the name cruciform corolla. Valvate aestivation.
Androecium: Stamens 6, polyandrous, arranged in two whorls of 4 and 2 (tetradynamous), outer two are short and the inner four are long, anthers bilobed, basifixed, introse.
Gynoecium: Bicarpellary, syncarpous, initially unilocular and later bilocular, (formation of pseudoseptum), one or more ovules on parietal placentation, style short, stigma bifid, sometimes bilobed, ovary superior.
Fruit: Siliqua or silicula.
Seeds: Endospermic
fig. 27.31a Habit, floral diagram and floral formula
Common Examples
Edible
1. Brassica oleracea Has many varieties
B.oleracea var.capitata - cabbage
B.oleracea var. botrytis - knol khol
B.oleracea var. caulorapa - corm
2. Raphanus sativus - radish
3. Brassica campestris - yellow mustard seeds yield oil
Medicinal
1. Sysimbrium officinale leaves and stem used in curing scurvy.
2. Nasturtium indica seeds are used for treating asthma.
3. Eruca sativa oil extracted from the plant used for treating burns.
4. Lepidium sativum seeds are used for treating liver disorders.
Ornamental
1. Iberis amara (candy tuft)
2. Cheiranthus sp.
fig. 27.32 Examples of Family Brassicaceae
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Angiosperm Families
Introduction
Family Poaceae (Graminae)
Family Liliaceae
Family Fabaceae
Sub Family Papillionatae (= Faboideae)
Sub Family: Caesalpiniodeae (Caesalpinae)
Sub Family: Mimosae (Mimosoidae)
Family Brassicaceae (= Cruciferae)
Family Solanaceae
Family Asteraceae
Subjects
English
Math
Geometry
Algebra
Number Theory
Discrete Mathematics
Boolean Algebra
Trigonometry
Statistics and Probability
Calculus
Physics
Science
Statistics
Biology
Chemistry
Saturday, January 31, 2009
solanaceae
Division: Angiospermae
Class: Dicotyledonae
Sub Class: Gamopetalae
Series: Bicarpellatae
Order: Tubiflorae
Family: Solanaceae
Solanaceae is commonly called as 'Brinjal family'. It includes about 90 genera and 2000 species. Members have a cosmopolitan distribution. There are around 60 species found in India.
Characteristic Features
Habit: Mostly annual herbs (Solanum nigrum), undershrubs (Solanum melongena). Some are shrubs (Solanum torvum) rarely trees (Solanum grandiflorum) and climbers (Solanum seaforthianum).
fig. 27.33 Habit and L.S. of flower of Solanum nigrum
fig. 27.34 Solanum nigrum
Root: Taproot, branched.
Stem: Herbaceous or woody, branched, often with prickles.
Leaves: Simple, alternate, sometimes pinnately divided into unequal lobes, exstipulate.
Inflorescence: Cymose. Often solitary cyme.
Flower: Ebracteate, actinomorphic, bisexual, pedicellate, heterochlamydeous, hypogynous.
Calyx: Sepals 5, gamosepalous, persistent (in the fruit condition also) valvate aestivation.
Corolla: Petals 5, gamopetalous, rotate or companulate or bilabiate (Schizanthus) or infundibuliform (Datura). Twisted or valvate aestivation.
fig. 27.35 Datura
Androecium: Stamens 5, epipetalous.
Gynoecium: Bicarpellary, syncarpous, bilocular or tetralocular due to pseudoseptum. Many ovules on axile placentation. Ovary superior, Obliquely placed.
Fruit: Berry or capsule.
Seed: Endospermic.
fig. 27.36 Floral parts, floral diagram and floral formula
Common Examples
Edible
1. Solanum tuberosum potato
2. Solanum melongena brinjal
3. Lycopersicum esculentum tomato
4. Capsicum annum red chilli
5. Physalis peruviana Raspberry.
Medicinal
1. Atropa belladona used for making belladona plasters. Atropine is a medicinal extract.
2. Hyoscyamus niger henbane used as sedative
3. Withania somnifera Ashwagandha used as an aphrodisiac.
4. Datura stromanium used as sedative and intoxicant.
Ornamental
1. Cestum nocturnum night queen or night jasmine
2. Petunia
3. Browalia
Others
1. Nicotiana tobaccum cured leaves used as tobacco, contains nicotine.
fig. 27.37 Examples of Family Solanaceae
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Angiosperm Families
Introduction
Family Poaceae (Graminae)
Family Liliaceae
Family Fabaceae
Sub Family Papillionatae (= Faboideae)
Sub Family: Caesalpiniodeae (Caesalpinae)
Sub Family: Mimosae (Mimosoidae)
Family Brassicaceae (= Cruciferae)
Family Solanaceae
Family Asteraceae
Subjects
English
Math
Geometry
Algebra
Number Theory
Discrete Mathematics
Boolean Algebra
Trigonometry
Statistics and Probability
Calculus
Physics
Science
Statistics
Biology
Chemistry
Class: Dicotyledonae
Sub Class: Gamopetalae
Series: Bicarpellatae
Order: Tubiflorae
Family: Solanaceae
Solanaceae is commonly called as 'Brinjal family'. It includes about 90 genera and 2000 species. Members have a cosmopolitan distribution. There are around 60 species found in India.
Characteristic Features
Habit: Mostly annual herbs (Solanum nigrum), undershrubs (Solanum melongena). Some are shrubs (Solanum torvum) rarely trees (Solanum grandiflorum) and climbers (Solanum seaforthianum).
fig. 27.33 Habit and L.S. of flower of Solanum nigrum
fig. 27.34 Solanum nigrum
Root: Taproot, branched.
Stem: Herbaceous or woody, branched, often with prickles.
Leaves: Simple, alternate, sometimes pinnately divided into unequal lobes, exstipulate.
Inflorescence: Cymose. Often solitary cyme.
Flower: Ebracteate, actinomorphic, bisexual, pedicellate, heterochlamydeous, hypogynous.
Calyx: Sepals 5, gamosepalous, persistent (in the fruit condition also) valvate aestivation.
Corolla: Petals 5, gamopetalous, rotate or companulate or bilabiate (Schizanthus) or infundibuliform (Datura). Twisted or valvate aestivation.
fig. 27.35 Datura
Androecium: Stamens 5, epipetalous.
Gynoecium: Bicarpellary, syncarpous, bilocular or tetralocular due to pseudoseptum. Many ovules on axile placentation. Ovary superior, Obliquely placed.
Fruit: Berry or capsule.
Seed: Endospermic.
fig. 27.36 Floral parts, floral diagram and floral formula
Common Examples
Edible
1. Solanum tuberosum potato
2. Solanum melongena brinjal
3. Lycopersicum esculentum tomato
4. Capsicum annum red chilli
5. Physalis peruviana Raspberry.
Medicinal
1. Atropa belladona used for making belladona plasters. Atropine is a medicinal extract.
2. Hyoscyamus niger henbane used as sedative
3. Withania somnifera Ashwagandha used as an aphrodisiac.
4. Datura stromanium used as sedative and intoxicant.
Ornamental
1. Cestum nocturnum night queen or night jasmine
2. Petunia
3. Browalia
Others
1. Nicotiana tobaccum cured leaves used as tobacco, contains nicotine.
fig. 27.37 Examples of Family Solanaceae
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Angiosperm Families
Introduction
Family Poaceae (Graminae)
Family Liliaceae
Family Fabaceae
Sub Family Papillionatae (= Faboideae)
Sub Family: Caesalpiniodeae (Caesalpinae)
Sub Family: Mimosae (Mimosoidae)
Family Brassicaceae (= Cruciferae)
Family Solanaceae
Family Asteraceae
Subjects
English
Math
Geometry
Algebra
Number Theory
Discrete Mathematics
Boolean Algebra
Trigonometry
Statistics and Probability
Calculus
Physics
Science
Statistics
Biology
Chemistry
papilionatae
perennial herbs and shrubs, trees are also common. Some are climbers (Pisum, Lathyrus).
fig. 27.17 Habit, L.S. of flower of Pisum sativum
fig. 27.18 Lathyrus and Pisum
Root: Branched taproot, usually with nodules containing bacteria of genus Rhizobium.
Stem: Herbaceous or woody, branched.
Leaf: Simple or compound, alternate, stipulate, leaf base is pulvinate.
Inflorescence: Racemose, simple or compound.
Flower: Bracteate, zygomorphic, bisexual, pedicellate, heterochlamydeous, pentamerous, hypogynous.
fig. 27.19 Flower
Calyx: Sepals 5, gamosepalous, hairy, valvate or imbricate aestivation.
Corolla: Petals 5, polypetalous, papillionaceous, with a standard (vexillum) petal, two wing petals (Alae) and two keel petals (carinae). Vexillary aestivation.
Androecium: Stamens 10, monadelphous or diadelphous, anthers dithecous.
fig.27.20 Androecium
Gynoecium: Monocarpellary, unilocular with ovules on marginal placentation. Ovary superior.
fig. 27.21 Gynoecium and TS of ovary
Fruit: A legume or lomentum.
Seed: Large, Non-endospermic
fig.27.22 L.S. of fruit and seeds
fig.27.23 Floral diagram
Floral formula
Common Examples
Pulses
1. Pisum sativum (pea)
2. Dolichos lablab (country bean)
3. Dolichos biflorus (horse gram)
4. Phaseolus mungo (black gram)
5. Phaseolus radiatus (green gram)
6. Cajanus cajan (red gram)
7. Glycine max (soya bean)
8. Cicer arientium (bengal gram)
Oil Yielding
1. Arachis hypogea (ground nut)
2. Pongamia pinnata (Pongam oil)
Timber Yielding
1. Dalbergia latifolia (Rosewood)
2. Dalbergia sisso (Sissoo)
Fibre Yielding
1. Crotolaria juncea (Sun hemp)
Medicinal
1. Clitoria ternatea used as antidote for snake bite in traditional practice of snake bite.
2. Desomodium gangeticus for curing cough and asthma.
Others
1. Indigofera tinctorea dye yielding (the famous of indigo of commerce)
2. Abrus precatorius Mature seeds sand used as weights by gold smiths in the past.
3. Butea monosperma ornamental plant, used for making eating plates.
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Angiosperm Families
Introduction
Family Poaceae (Graminae)
Family Liliaceae
Family Fabaceae
Sub Family Papillionatae (= Faboideae)
Sub Family: Caesalpiniodeae (Caesalpinae)
Sub Family: Mimosae (Mimosoidae)
Family Brassicaceae (= Cruciferae)
Family Solanaceae
Family Asteraceae
SubjectsEnglishMathGeometry
AlgebraNumber TheoryDiscrete Mathematics
Boolean AlgebraTrigonometryStatistics and Probability
CalculusPhysicsScience
StatisticsBiologyChemistry
Home About Us Press Room Contact Us Teaching Jobs Privacy Policy Help Site Map Terms Of Use Essay Grading Library Programs
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fig. 27.17 Habit, L.S. of flower of Pisum sativum
fig. 27.18 Lathyrus and Pisum
Root: Branched taproot, usually with nodules containing bacteria of genus Rhizobium.
Stem: Herbaceous or woody, branched.
Leaf: Simple or compound, alternate, stipulate, leaf base is pulvinate.
Inflorescence: Racemose, simple or compound.
Flower: Bracteate, zygomorphic, bisexual, pedicellate, heterochlamydeous, pentamerous, hypogynous.
fig. 27.19 Flower
Calyx: Sepals 5, gamosepalous, hairy, valvate or imbricate aestivation.
Corolla: Petals 5, polypetalous, papillionaceous, with a standard (vexillum) petal, two wing petals (Alae) and two keel petals (carinae). Vexillary aestivation.
Androecium: Stamens 10, monadelphous or diadelphous, anthers dithecous.
fig.27.20 Androecium
Gynoecium: Monocarpellary, unilocular with ovules on marginal placentation. Ovary superior.
fig. 27.21 Gynoecium and TS of ovary
Fruit: A legume or lomentum.
Seed: Large, Non-endospermic
fig.27.22 L.S. of fruit and seeds
fig.27.23 Floral diagram
Floral formula
Common Examples
Pulses
1. Pisum sativum (pea)
2. Dolichos lablab (country bean)
3. Dolichos biflorus (horse gram)
4. Phaseolus mungo (black gram)
5. Phaseolus radiatus (green gram)
6. Cajanus cajan (red gram)
7. Glycine max (soya bean)
8. Cicer arientium (bengal gram)
Oil Yielding
1. Arachis hypogea (ground nut)
2. Pongamia pinnata (Pongam oil)
Timber Yielding
1. Dalbergia latifolia (Rosewood)
2. Dalbergia sisso (Sissoo)
Fibre Yielding
1. Crotolaria juncea (Sun hemp)
Medicinal
1. Clitoria ternatea used as antidote for snake bite in traditional practice of snake bite.
2. Desomodium gangeticus for curing cough and asthma.
Others
1. Indigofera tinctorea dye yielding (the famous of indigo of commerce)
2. Abrus precatorius Mature seeds sand used as weights by gold smiths in the past.
3. Butea monosperma ornamental plant, used for making eating plates.
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Angiosperm Families
Introduction
Family Poaceae (Graminae)
Family Liliaceae
Family Fabaceae
Sub Family Papillionatae (= Faboideae)
Sub Family: Caesalpiniodeae (Caesalpinae)
Sub Family: Mimosae (Mimosoidae)
Family Brassicaceae (= Cruciferae)
Family Solanaceae
Family Asteraceae
SubjectsEnglishMathGeometry
AlgebraNumber TheoryDiscrete Mathematics
Boolean AlgebraTrigonometryStatistics and Probability
CalculusPhysicsScience
StatisticsBiologyChemistry
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fabaceae
Systematic Position
Division: Angiospermae
Class: Dicotyledonae
Sub Class: Polypetalae
Series: Calyciflorae
Order: Rosales
Family: Fabaceae (=Leguminosae)
Leguminosae is one of the largest families in dicots, including about 690 genera and nearly 18,000 species. In all the major systems of classification, the family Leguminosae has been traditionally considered to include three sub families Papillionaceae, Caesalpinae and Mimosae. On the basis of vegetative characters, it is very difficult to characterise the sub families. However, the following features, with some exception, can be considered as common to all the three subfamilies of Leguminosae.
Roots contain root nodules that harbour symbiotic, nitrogen fixing bacteria.
Leaves are compound, alternate, stipulate and pulvinate.
Inflorescence is variable form of racemose types.
Flowers are generally zygomorphic, hypogynous, heterochlamydeous, pentamerous and hypogynous.
Stamens are usually ten in number, monadelphous or diadelphous or free.
Gynoecium is monocarpellary, unilocular with marginal placentation.
Fruit is a Legume or lomentum.
Division: Angiospermae
Class: Dicotyledonae
Sub Class: Polypetalae
Series: Calyciflorae
Order: Rosales
Family: Fabaceae (=Leguminosae)
Leguminosae is one of the largest families in dicots, including about 690 genera and nearly 18,000 species. In all the major systems of classification, the family Leguminosae has been traditionally considered to include three sub families Papillionaceae, Caesalpinae and Mimosae. On the basis of vegetative characters, it is very difficult to characterise the sub families. However, the following features, with some exception, can be considered as common to all the three subfamilies of Leguminosae.
Roots contain root nodules that harbour symbiotic, nitrogen fixing bacteria.
Leaves are compound, alternate, stipulate and pulvinate.
Inflorescence is variable form of racemose types.
Flowers are generally zygomorphic, hypogynous, heterochlamydeous, pentamerous and hypogynous.
Stamens are usually ten in number, monadelphous or diadelphous or free.
Gynoecium is monocarpellary, unilocular with marginal placentation.
Fruit is a Legume or lomentum.
graminae
Family Poaceae (Graminae)
Systematic Position
Division: Angiosperms
Class: Monocotyledonae
Family: Poaceae (Graminae) (The grass family)
It is one of the largest families in monocots consisting of 620 genera and about 6000 species. Members are cosmopolitan in distribution. Around 900 species are present in India.
Characteristic Features
Habit: Mainly herbs (annuals or perennials) or shrubs. Some are trees like (Bambusa, Dendrocalamus).
fig. 27.2 Bambusa and Dendrocalamus
Root: Adventitious, fibrous, branched or stilt (as in maize).
Stem: Underground rhizome in all perennial grasses, cylindrical, distinct nodes and internodes, herbaceous or woody.
Leaves: Alternate, simple, extipulate, sessile, leaf base forming tubular sheath, sheath open, surrounding the internodes completely, hairy or rough, linear, parallel venation.
Inflorescence: Compound spike, sessile or stalked. Each unit is called spikelet, may be a spike of spikelets (Triticum) or panicle of spikelets (Avena).
fig. 27.3 Avena and Triticum
Flower: Bracteate, bracteolate, sessile, incomplete, bisexual or unisexual (Zea mays), zygomorphic, hypogynous, homochlamydeous.
fig. 27.4 Zea mays
Perianth: Represented by membranous scales called lodicules, many (Ochlandra) or three or two or absent.
fig. 27.5
Androecium: Stamens usually three, some times six (Bambusa) rarely one (species of Fistuca). Filaments long, anthers dithecous, versatile and linear.
Gynoecium: Monocarpellary (presumed to be three of which two are aborted), unilocular, single ovule on basal placentation, style short or absent, stigma bifid, ovary superior.
Fruit: A caryopsis with pericarp completely united with the seed coat, rarely a nut (Dendrocalamus) or a berry (Bambusa).
Seed: Endospermic, with a single cotyledon called scutellum, pressed against the endosperm
Floral diagram and formula
fig. 27.6 Floral diagram and formula of male flower
fig.27.7 Floral diagram and formula of female flower
Common Examples
Cereals
1. Triticum aestivum (wheat)
2. Zea mays (Maize)
3. Oryza sativa (Rice)
4. Hordeum vulgare (Barley)
5. Avena sativum (Oat)
fig. 27.8 Examples of Cereals
Millets
1. Sorghum vulgare (Jawar)
2. Pennisetum typhoides (Bajra)
fig. 27.9 Examples of Millets
Grasses
1. Cynodon dactylon (Dog grass)
2. Andropogon muricatus (Khas grass-oil yielding)
fig. 27.10 Examples of Grasses
Paper Grasses
1. Themeda gigantia (Kapoor grass)
2. Eucaliopsis binala (Bhabar grass)
fig. 27.11 Bhabar Grass
Other
1. Bambusa vulgans. (Bamboo plant of commercial ornamental importance)
2. Saccharum officinarum (Sugarcane plant)
fig. 27.12 Bamboo and Sugarcane Plants
Systematic Position
Division: Angiosperms
Class: Monocotyledonae
Family: Poaceae (Graminae) (The grass family)
It is one of the largest families in monocots consisting of 620 genera and about 6000 species. Members are cosmopolitan in distribution. Around 900 species are present in India.
Characteristic Features
Habit: Mainly herbs (annuals or perennials) or shrubs. Some are trees like (Bambusa, Dendrocalamus).
fig. 27.2 Bambusa and Dendrocalamus
Root: Adventitious, fibrous, branched or stilt (as in maize).
Stem: Underground rhizome in all perennial grasses, cylindrical, distinct nodes and internodes, herbaceous or woody.
Leaves: Alternate, simple, extipulate, sessile, leaf base forming tubular sheath, sheath open, surrounding the internodes completely, hairy or rough, linear, parallel venation.
Inflorescence: Compound spike, sessile or stalked. Each unit is called spikelet, may be a spike of spikelets (Triticum) or panicle of spikelets (Avena).
fig. 27.3 Avena and Triticum
Flower: Bracteate, bracteolate, sessile, incomplete, bisexual or unisexual (Zea mays), zygomorphic, hypogynous, homochlamydeous.
fig. 27.4 Zea mays
Perianth: Represented by membranous scales called lodicules, many (Ochlandra) or three or two or absent.
fig. 27.5
Androecium: Stamens usually three, some times six (Bambusa) rarely one (species of Fistuca). Filaments long, anthers dithecous, versatile and linear.
Gynoecium: Monocarpellary (presumed to be three of which two are aborted), unilocular, single ovule on basal placentation, style short or absent, stigma bifid, ovary superior.
Fruit: A caryopsis with pericarp completely united with the seed coat, rarely a nut (Dendrocalamus) or a berry (Bambusa).
Seed: Endospermic, with a single cotyledon called scutellum, pressed against the endosperm
Floral diagram and formula
fig. 27.6 Floral diagram and formula of male flower
fig.27.7 Floral diagram and formula of female flower
Common Examples
Cereals
1. Triticum aestivum (wheat)
2. Zea mays (Maize)
3. Oryza sativa (Rice)
4. Hordeum vulgare (Barley)
5. Avena sativum (Oat)
fig. 27.8 Examples of Cereals
Millets
1. Sorghum vulgare (Jawar)
2. Pennisetum typhoides (Bajra)
fig. 27.9 Examples of Millets
Grasses
1. Cynodon dactylon (Dog grass)
2. Andropogon muricatus (Khas grass-oil yielding)
fig. 27.10 Examples of Grasses
Paper Grasses
1. Themeda gigantia (Kapoor grass)
2. Eucaliopsis binala (Bhabar grass)
fig. 27.11 Bhabar Grass
Other
1. Bambusa vulgans. (Bamboo plant of commercial ornamental importance)
2. Saccharum officinarum (Sugarcane plant)
fig. 27.12 Bamboo and Sugarcane Plants
In the description of Angiosperms, family is the most useful taxonomic unit. Ranks higher than family are not of much practical significance and are used mostly in classification systems and academic discussions. The taxonomic category genus also does not get much importance as plants are scientifically described in binomial nomenclature. Use of only the generic name isolated from the specific name may lead to confusion. The genus Solanum includes several species of which Solanum tuberosum alone represents the potato plant.
Family Poaceae (Graminae)
It is one of the largest families in monocots consisting of 620 genera and about 6000 species. Members are cosmopolitan in distribution. Around 900 species are present in India.
Family Liliaceae
It is commonly called Lily family. It includes about 250 genera and 3700 species that have a cosmopolitan distribution. Around 200 species are available in India.
Family Fabaceae
Leguminosae is one of the largest families in dicots, including about 690 genera and nearly 18,000 species. In all the major systems of classification, the family Leguminosae has been traditionally considered to include three sub families Papillionaceae, Caesalpinae and Mimosae. On the basis of vegetative characters, it is very difficult to characterise the sub families. However, the following features, with some exception, can be considered as common to all the three subfamilies of Leguminosae.
Sub Family Papillionatae (= Faboideae)
Habit: Mostly perennial herbs and shrubs, trees are also common. Some are climbers (Pisum, Lathyrus).
Sub Family: Caesalpiniodeae (Caesalpinae)
Habit: Mostly trees and shrubs, rarely herbs, some are climbers (Bauhinia).
Sub Family: Mimosae (Mimosoidae)
Habit: Herbs (Mimosa), shrubs and trees (Acacia).
Family Brassicaceae (= Cruciferae)
This family is commonly called as 'mustard family'. It includes about 300 genus and about 3700 species. The members have a cosmopolitan distribution. There are around 150 species in India.
Family Solanaceae
Solanaceae is commonly called as 'Brinjal family'. It includes about 90 genera and 2000 species. Members have a cosmopolitan distribution. There are around 60 species found in India.
Family Asteraceae
Commonly known as sunflower family. It is the largest family in dicots. It includes about 900 genera and 13,000 species. It has a cosmopolitan distribution. There are around 100 species in India.
Family Poaceae (Graminae)
It is one of the largest families in monocots consisting of 620 genera and about 6000 species. Members are cosmopolitan in distribution. Around 900 species are present in India.
Family Liliaceae
It is commonly called Lily family. It includes about 250 genera and 3700 species that have a cosmopolitan distribution. Around 200 species are available in India.
Family Fabaceae
Leguminosae is one of the largest families in dicots, including about 690 genera and nearly 18,000 species. In all the major systems of classification, the family Leguminosae has been traditionally considered to include three sub families Papillionaceae, Caesalpinae and Mimosae. On the basis of vegetative characters, it is very difficult to characterise the sub families. However, the following features, with some exception, can be considered as common to all the three subfamilies of Leguminosae.
Sub Family Papillionatae (= Faboideae)
Habit: Mostly perennial herbs and shrubs, trees are also common. Some are climbers (Pisum, Lathyrus).
Sub Family: Caesalpiniodeae (Caesalpinae)
Habit: Mostly trees and shrubs, rarely herbs, some are climbers (Bauhinia).
Sub Family: Mimosae (Mimosoidae)
Habit: Herbs (Mimosa), shrubs and trees (Acacia).
Family Brassicaceae (= Cruciferae)
This family is commonly called as 'mustard family'. It includes about 300 genus and about 3700 species. The members have a cosmopolitan distribution. There are around 150 species in India.
Family Solanaceae
Solanaceae is commonly called as 'Brinjal family'. It includes about 90 genera and 2000 species. Members have a cosmopolitan distribution. There are around 60 species found in India.
Family Asteraceae
Commonly known as sunflower family. It is the largest family in dicots. It includes about 900 genera and 13,000 species. It has a cosmopolitan distribution. There are around 100 species in India.
reptiles zx
This listing is provided as a supplement to the article "The ABC's of Animal Taxonomy." It is intended as an illustration of the modern use of traditional, or Linnaean, taxonomy, and is NOT guaranteed to be totally accurate. (In point of fact, NO taxonomy can ever be guaranteed to be totally accurate. Taxonomy involves many judgement calls, and it is highly unlikely that any two authorities on a particular subject will ever agree on all the details of any taxonomic scheme.) The scheme presented was prepared by Peter Uetz, originally with the European Molecular Biology Laboratory in Heidelberg, Germany, and now with the University of Washington. Some modifications were made by dlb.
I do not intend to present an equivalent cladistic taxonomy of Class Reptilia, for two reasons. First, I don't know where one can be obtained, or even if the information necessary to create one has yet been acquired for the whole class; and, second, this newsletter isn't big enough to hold one if it does exist!
-- dlb
Subclass Anapsida (Note 1)
Order Testudines (turtles)
Suborder Cryptodira
Superfamily Testudinoidea
Family Chelydridae (Snapping Turtles)
Family Emydidae (Pond Turtles/Box and Water Turtles)
Family Testudinidae (Tortoises)
Superfamily Trionychoidea
Family Dermatemydidae (River Turtles)
Family Kinosternidae (Mud and Musk Turtles)
Family Carettochelyidae (Pig-nose Turtles)
Family Trionychidae (Softshell Turtles)
Superfamily Chelonioidea
Family Cheloniidae (Sea Turtles)
Family Dermochelyidae (Leatherback Turtles)
Suborder Pleurodira
Family Pelomedusidae (Afro-American Sideneck Turtles)
Family Chelidae (Austro-American Sideneck Turtles)
Subclass Lepidosauria
Order Rhynchocephalia
Suborder Sphenodontida
Family Sphenodontidae (Tuataras)
Order Squamata ("scaled reptiles")
Suborder Sauria (lizards)
Infraorder Iguania
Family Agamidae (Agamas)
Family Chamaeleonidae (Chameleons)
Family Iguanidae (Iguanas and Spinytail Iguanas)
Family Corytophanidae (Casquehead Lizards)
Family Crotaphytidae (Collared and Leopard Lizards)
Family Hoplocercidae
Family Opluridae (Madagascar iguanids)
Family Phrynosomatidae (Earless, Spiny, Tree, Side-blotched and Horned Lizards)
Family Polychrotidae (Anoles)
Family Tropiduridae (Neotropical Ground Lizards)
Infraorder Gekkota
Family Gekkonidae (Geckoes)
Family Eublepharidae (Eyelid Geckoes)
Family Pygopodidae (Legless Lizards)
Family Dibamidae (Blind Lizards)
Infraorder Scincomorpha
Family Cordylidae (Spinytail Lizards)
Family Gerrhosauridae
Family Gymnophthalmidae (Spectacled Lizards)
Family Teiidae (Whiptails and Tegus)
Family Lacertidae (Wall Lizards)
Family Scincidae (Skinks)
Family Xantusiidae (Night Lizards)
Infraorder Diploglossa
Family Anguidae (Glass Lizards and Alligator Lizards; Lateral Fold Lizards)
Family Anniellidae (American Legless lizards)
Family Xenosauridae (Knob-scaled Lizards)
Infraorder Platynota (Varanoidea)
Family Helodermatidae (Gila Monsters)
Family Lanthanotidae (Earless Monitor lizards)
Family Varanidae (Monitor Lizards)
Suborder Serpentes (snakes)
Superfamily Typhlopoidea (Scolecophidia)
Family Anomalepidae (Dawn Blind Snakes)
Family Typhlopidae (Blind Snakes)
Family Leptotyphlopidae/Glauconiidae (Slender Blind Snakes)
Superfamily Henophidia (Boidea)
Family Aniliidae/Ilysiidae (Pipe Snakes)
Family Anomochilidae (Dwarf Pipe Snakes)
Family Boidae (Boas and Pythons)
Family Bolyeridae (Round Island Boas)
Family Cylindrophiidae (Asian Pipe Snakes)
Family Loxocemidae (Mexican Burrowing Pythons)
Family Tropidophiidae (Woodsnakes, Round Island "Boas ")
Family Uropeltidae (Short-tail Snakes)
Family Xenopeltidae (Sunbeam Snakes)
Superfamily Xenophidia (Colubroidea = Caenophidia)
Family Acrochordidae (File Snakes)
Family Atractaspididae (Mole Vipers)
Family Colubridae (Colubrids)
Family Elapidae (Cobras, Kraits, Coral Snakes)
Family Hydrophiidae (Sea Snakes)
Family Viperidae (Vipers and Pit Vipers)
Suborder Amphisbaenia (amphisbaenians)
Family Amphisbaenidae (Worm Lizards)
Family Trogonophidae (Shorthead Worm Lizards)
Family Bipedidae (Two-legged Worm Lizards)
Subclass Archosauria
Order Crocodylia
Suborder Eusuchia
Family Crocodylidae (Crocodiles)
Note 1: The taxonomic position of turtles is quite controversial at the present time. This classification represents the traditional, conservative view. (Return)Source: Modified from The EMBL Reptile Database, EMBL (European Molecular Biology Laboratory), Heidelberg, Germany. (Maintained by Peter Uetz, University of Washington. Modified by dlb) Taken off the Web, at <http://www.embl-heidelberg.de/~uetz/LivingReptiles.html>.
I do not intend to present an equivalent cladistic taxonomy of Class Reptilia, for two reasons. First, I don't know where one can be obtained, or even if the information necessary to create one has yet been acquired for the whole class; and, second, this newsletter isn't big enough to hold one if it does exist!
-- dlb
Subclass Anapsida (Note 1)
Order Testudines (turtles)
Suborder Cryptodira
Superfamily Testudinoidea
Family Chelydridae (Snapping Turtles)
Family Emydidae (Pond Turtles/Box and Water Turtles)
Family Testudinidae (Tortoises)
Superfamily Trionychoidea
Family Dermatemydidae (River Turtles)
Family Kinosternidae (Mud and Musk Turtles)
Family Carettochelyidae (Pig-nose Turtles)
Family Trionychidae (Softshell Turtles)
Superfamily Chelonioidea
Family Cheloniidae (Sea Turtles)
Family Dermochelyidae (Leatherback Turtles)
Suborder Pleurodira
Family Pelomedusidae (Afro-American Sideneck Turtles)
Family Chelidae (Austro-American Sideneck Turtles)
Subclass Lepidosauria
Order Rhynchocephalia
Suborder Sphenodontida
Family Sphenodontidae (Tuataras)
Order Squamata ("scaled reptiles")
Suborder Sauria (lizards)
Infraorder Iguania
Family Agamidae (Agamas)
Family Chamaeleonidae (Chameleons)
Family Iguanidae (Iguanas and Spinytail Iguanas)
Family Corytophanidae (Casquehead Lizards)
Family Crotaphytidae (Collared and Leopard Lizards)
Family Hoplocercidae
Family Opluridae (Madagascar iguanids)
Family Phrynosomatidae (Earless, Spiny, Tree, Side-blotched and Horned Lizards)
Family Polychrotidae (Anoles)
Family Tropiduridae (Neotropical Ground Lizards)
Infraorder Gekkota
Family Gekkonidae (Geckoes)
Family Eublepharidae (Eyelid Geckoes)
Family Pygopodidae (Legless Lizards)
Family Dibamidae (Blind Lizards)
Infraorder Scincomorpha
Family Cordylidae (Spinytail Lizards)
Family Gerrhosauridae
Family Gymnophthalmidae (Spectacled Lizards)
Family Teiidae (Whiptails and Tegus)
Family Lacertidae (Wall Lizards)
Family Scincidae (Skinks)
Family Xantusiidae (Night Lizards)
Infraorder Diploglossa
Family Anguidae (Glass Lizards and Alligator Lizards; Lateral Fold Lizards)
Family Anniellidae (American Legless lizards)
Family Xenosauridae (Knob-scaled Lizards)
Infraorder Platynota (Varanoidea)
Family Helodermatidae (Gila Monsters)
Family Lanthanotidae (Earless Monitor lizards)
Family Varanidae (Monitor Lizards)
Suborder Serpentes (snakes)
Superfamily Typhlopoidea (Scolecophidia)
Family Anomalepidae (Dawn Blind Snakes)
Family Typhlopidae (Blind Snakes)
Family Leptotyphlopidae/Glauconiidae (Slender Blind Snakes)
Superfamily Henophidia (Boidea)
Family Aniliidae/Ilysiidae (Pipe Snakes)
Family Anomochilidae (Dwarf Pipe Snakes)
Family Boidae (Boas and Pythons)
Family Bolyeridae (Round Island Boas)
Family Cylindrophiidae (Asian Pipe Snakes)
Family Loxocemidae (Mexican Burrowing Pythons)
Family Tropidophiidae (Woodsnakes, Round Island "Boas ")
Family Uropeltidae (Short-tail Snakes)
Family Xenopeltidae (Sunbeam Snakes)
Superfamily Xenophidia (Colubroidea = Caenophidia)
Family Acrochordidae (File Snakes)
Family Atractaspididae (Mole Vipers)
Family Colubridae (Colubrids)
Family Elapidae (Cobras, Kraits, Coral Snakes)
Family Hydrophiidae (Sea Snakes)
Family Viperidae (Vipers and Pit Vipers)
Suborder Amphisbaenia (amphisbaenians)
Family Amphisbaenidae (Worm Lizards)
Family Trogonophidae (Shorthead Worm Lizards)
Family Bipedidae (Two-legged Worm Lizards)
Subclass Archosauria
Order Crocodylia
Suborder Eusuchia
Family Crocodylidae (Crocodiles)
Note 1: The taxonomic position of turtles is quite controversial at the present time. This classification represents the traditional, conservative view. (Return)Source: Modified from The EMBL Reptile Database, EMBL (European Molecular Biology Laboratory), Heidelberg, Germany. (Maintained by Peter Uetz, University of Washington. Modified by dlb) Taken off the Web, at <http://www.embl-heidelberg.de/~uetz/LivingReptiles.html>.
animal taxonomy
There are two distinct taxonomic systems currently in vogue among professional zoologists today. The traditional, or Linnaean, taxonomy is still largely in favor among field workers, conservationists, and husbandry people. The alternative, Cladistic taxonomy, is overwhelmingly supported by evolutionary biologists.
The undisputed father of taxonomy was the Swedish botanist Karl von Linné (1707-1778). Because virtually all science in the 18th century was written in Latin, von Linné is better known by his Latinized name, Carolus Linnaeus. Linnaeus created the system of scientific nomenclature still in use today, wherein every species is given two Latin names, a genus, or group name, and the species name. In formal Latin usage, the genus name is a noun and is capitalized, while the species name is an adjective, which should agree in gender with the genus, and is written in lower case.
Over time, biologist added additional, larger and higher level group names, called taxons (plural: taxa), from Family up to Kingdom, arranged in a hierarchical order, until a standardized 7-level hierarchy was established, as follows:
Kingdom
Animalia
Phylum
Chordata
Class
Reptilia
Order
Squamata
Family
Colubridae
Genus
Pituophis
species
catenifer
(Bullsnake, Gopher Snake)
To further facilitate grouping similar or closely related groups, these taxa may optionally be divided with from one to three named intermediate-level taxa, as required. For example:
Class
Major division (required)
Subclass
1st optional subdivision
Infraclass
3rd optional subdivision
Superorder
2nd optional subdivision
Order
Major division (required)The taxa Superkingdom and Infraspecies are generally not used, leaving a maximum of 26 possible taxonomic categories, although all are rarely required for any given species (or subspecies).
One sub-taxon that is most commonly used is the subspecies, as in Pituophis catenifer sayi, the bullsnake, as shown in the example above. Others that should be familiar to herpers are Suborder Lacertilia (lizards), and Suborder Serpentes (snakes), two divisions of the Order Squamata. (The third division of Squamata is Suborder Amphisbaenia, the worm lizards.)
The original purpose of taxonomy was the recognition, categorization, and identification of organisms. Therefore, species were grouped into higher level taxa based primarily by apparent resemblance or by the possession of shared traits. With the widespread acceptance of the theory of evolution, more of an attempt was made to group species in accordance with their evolutionary history. This endeavor is what lead to the creation of the sub-taxon groups, in order that taxonomists could more accurately depict evolutionary relationships between species or other taxa. As our understanding of evolution increased, and our tools for determining evolutionary history and genetic relationships became more sophisticated, the task of ordering species into only 26 or 28 taxons became impossible. That is when cladistics was invented.
A clade is just a fancy word for group, but in cladistic taxonomy, it has a very precise definition. Any clade can be defined as two member species (or any other taxonomic group) and all the descendants of their nearest common ancestor. Depicted graphically as a cladogram, relationships are shown on a binary tree, where every fork has exactly two branches, each clade represented by a fork. No matter what the relationship is between the groups depicted in a cladogram, there will always be one less clade than the number of groups. Thus, any cladogram depicting eight taxonomic groups will always contain seven clades, regardless of their arrangement, as shown:
Cladogram 1.Groups G and H are most closely related, while group A is the most distant. Clades are represented by numbers 1 through 7.
Cladogram 2.Groups A and B are more closely related to each other than to any other group, as are C and D, E and F, and G and H. Clade 4 is more closely related to 5, and clade 6 is more closely related to 7.
Due to the sheer number of clades (all of which are generally assigned names), cladistic taxonomy becomes impractical when trying to simultaneously describe all of the world's 3,000 some odd reptile species, let alone the two to ten million species of plants, animals, and micro-organisms that are known or presumed to exist. It does, however, provide a very powerful tool for analyzing relationships within smaller assemblages of organisms.
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The undisputed father of taxonomy was the Swedish botanist Karl von Linné (1707-1778). Because virtually all science in the 18th century was written in Latin, von Linné is better known by his Latinized name, Carolus Linnaeus. Linnaeus created the system of scientific nomenclature still in use today, wherein every species is given two Latin names, a genus, or group name, and the species name. In formal Latin usage, the genus name is a noun and is capitalized, while the species name is an adjective, which should agree in gender with the genus, and is written in lower case.
Over time, biologist added additional, larger and higher level group names, called taxons (plural: taxa), from Family up to Kingdom, arranged in a hierarchical order, until a standardized 7-level hierarchy was established, as follows:
Kingdom
Animalia
Phylum
Chordata
Class
Reptilia
Order
Squamata
Family
Colubridae
Genus
Pituophis
species
catenifer
(Bullsnake, Gopher Snake)
To further facilitate grouping similar or closely related groups, these taxa may optionally be divided with from one to three named intermediate-level taxa, as required. For example:
Class
Major division (required)
Subclass
1st optional subdivision
Infraclass
3rd optional subdivision
Superorder
2nd optional subdivision
Order
Major division (required)The taxa Superkingdom and Infraspecies are generally not used, leaving a maximum of 26 possible taxonomic categories, although all are rarely required for any given species (or subspecies).
One sub-taxon that is most commonly used is the subspecies, as in Pituophis catenifer sayi, the bullsnake, as shown in the example above. Others that should be familiar to herpers are Suborder Lacertilia (lizards), and Suborder Serpentes (snakes), two divisions of the Order Squamata. (The third division of Squamata is Suborder Amphisbaenia, the worm lizards.)
The original purpose of taxonomy was the recognition, categorization, and identification of organisms. Therefore, species were grouped into higher level taxa based primarily by apparent resemblance or by the possession of shared traits. With the widespread acceptance of the theory of evolution, more of an attempt was made to group species in accordance with their evolutionary history. This endeavor is what lead to the creation of the sub-taxon groups, in order that taxonomists could more accurately depict evolutionary relationships between species or other taxa. As our understanding of evolution increased, and our tools for determining evolutionary history and genetic relationships became more sophisticated, the task of ordering species into only 26 or 28 taxons became impossible. That is when cladistics was invented.
A clade is just a fancy word for group, but in cladistic taxonomy, it has a very precise definition. Any clade can be defined as two member species (or any other taxonomic group) and all the descendants of their nearest common ancestor. Depicted graphically as a cladogram, relationships are shown on a binary tree, where every fork has exactly two branches, each clade represented by a fork. No matter what the relationship is between the groups depicted in a cladogram, there will always be one less clade than the number of groups. Thus, any cladogram depicting eight taxonomic groups will always contain seven clades, regardless of their arrangement, as shown:
Cladogram 1.Groups G and H are most closely related, while group A is the most distant. Clades are represented by numbers 1 through 7.
Cladogram 2.Groups A and B are more closely related to each other than to any other group, as are C and D, E and F, and G and H. Clade 4 is more closely related to 5, and clade 6 is more closely related to 7.
Due to the sheer number of clades (all of which are generally assigned names), cladistic taxonomy becomes impractical when trying to simultaneously describe all of the world's 3,000 some odd reptile species, let alone the two to ten million species of plants, animals, and micro-organisms that are known or presumed to exist. It does, however, provide a very powerful tool for analyzing relationships within smaller assemblages of organisms.
About Me
Web Site Design
Plate TectonicsPaleoclimatePaleontology
EvolutionTaxonomy
Herpetology
WebSpinners.com
Site designed by: WebSpinners (info@webspinners.com)Low Cost Web Design and Hosting ServiceWebmaster: Donald L. Blanchard
copied text plant physiology
Department of Biology, Pennsylvania State University, University Park, Pennsylvania 16802
Plants have specialized organs for distinct functions. Leaves perform photosynthesis and fix carbon, whereas roots absorb water and minerals. To distribute resources between these organs, plants have a vasculature composed of phloem and xylem. The xylem conducts water and minerals from the roots up to the shoots. The phloem transports carbon- and nitrogen-containing compounds from mature leaves to the roots and to other nonphotosynthetic organs such as flowers and fruits. Phloem tissue comprises two main cell types: sieve elements and companion cells. Sieve elements conduct nutrients, while companion cells metabolically support the sieve elements (van Bel and Knoblauch, 2000). The vascular system represents a highly integrated distribution network essential not only for the life of the plant but also for the life of the planet, as nearly all terrestrially produced chemical energy, including our food supply, is derived from plants.
Photosynthesis and carbon assimilation occur in leaf mesophyll cells and additionally in bundle sheath cells in C4 plants. For distribution to distal tissues, fixed carbon must move out of the photosynthetic cells and into the phloem. If the photoassimilates (assimilated carbon) diffuse down a concentration gradient from the mesophyll cells into the phloem following an entirely cytoplasmic path through plasmodesmata (intercellular channels through which small molecules freely diffuse), it is referred to as symplastic phloem loading (Turgeon and Medville, 1998). However, if the assimilated carbon must cross a membrane prior to entering into the phloem, it is called apoplastic phloem loading (van Bel, 1993; Turgeon, 2006). In this case, the concentration of the transported assimilate is higher in the phloem than in the photosynthetic cells. Because apoplastic phloem loading involves movement against a concentration gradient, it requires energy in the form of a pH gradient generated by H+-ATPases (Bush, 1993; Gaxiola et al., 2007).
Carbon partitioning is the process whereby assimilates are distributed throughout the plant body from photosynthetic tissues. For most plants, this occurs by loading Suc into the phloem and transporting it from source tissues (net exporters) to sink tissues (net importers), where Suc is unloaded (Turgeon, 1989; van Bel, 2003). This process is well characterized at the physiological, biochemical, and anatomical levels (Hofstra and Nelson, 1969; Fellows and Geiger, 1974; Evert et al., 1978; Nguyen-Quoc et al., 1990; Huber and Hanson, 1992; Evert et al., 1996a; Koch, 1996; Paul and Foyer, 2001). However, despite the obvious importance of this process for plant growth and development, few genes that function in carbon partitioning have been identified. Suc, K+, and water transporters/channels have been characterized for their contribution to the transport of Suc in the phloem (Deeken et al., 2002; Lalonde et al., 2004; Maurel, 2007; Sauer, 2007). Of these, the best described genes that directly control Suc loading into the phloem encode Suc transporters (SUTs; Lalonde et al., 2004; Sauer, 2007).
In this review, we discuss phloem loading and the control of carbon partitioning in grasses, focusing on SUTs and highlighting similarities and differences with eudicots. Additionally, we cover related aspects of phloem loading, such as leaf anatomy, and discuss other genes regulating carbohydrate accumulation in grass leaves. For additional discussions of genes that function in phloem loading and carbon partitioning (e.g. H+-ATPase, K+ channel, Suc synthase, aquaporins), we refer interested readers to the following articles (Nolte and Koch, 1993; DeWitt and Sussman, 1995; Hannah et al., 2001; Deeken et al., 2002; Dinges et al., 2003; Ma et al., 2004; Hardin et al., 2006; Lu and Sharkey, 2006; Smith and Stitt, 2007).
LEAF ANATOMY
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED As the path of phloem loading is intimately related to leaf structure, we begin with a brief overview of grass leaf anatomy, describing a maize (Zea mays) leaf blade as a typical example (Fig. 1A ). Three orders of veins are arranged longitudinally along the main axis of the leaf (Esau, 1977). The largest type contains large metaxylem vessels and a protoxylem lacuna (space; Fig. 1A). Additionally, as structural support, hypodermal sclerenchyma girders are located between the vein and the epidermal cell layers. Intermediate veins lack metaxylem vessels but have hypodermal sclerenchyma caps on one or both sides of the vein. The smallest veins have neither metaxylem vessels nor hypodermal sclerenchyma caps. Collectively, the intermediate and small veins are referred to as minor veins, while the largest veins are known as lateral or major veins. Minor veins intergrade into the lateral veins (Russell and Evert, 1985). Transverse veins with anatomy similar to the small veins connect adjacent longitudinal veins. All veins contain phloem cells that transport photoassimilates, but the various orders of veins play different roles in carbon partitioning. Minor veins are the site where photoassimilates are loaded into the vein (Fritz et al., 1983). Lateral veins principally function in the long-distance transport of assimilated carbon out of the leaf into the stem (Fritz et al., 1989). Transverse veins shunt assimilates from minor veins to lateral veins (Fritz et al., 1989).
View larger version (49K):[in this window][in a new window][as a PowerPoint slide]
Figure 1. Maize leaf anatomy and path of Suc movement. A, Cross section of a maize leaf blade viewed under UV light to visualize cellular anatomy. Chlorophyll autofluoresces red and cell walls autofluoresce blue. One lateral vein and three minor veins are shown. B, Schematic diagram of cell types associated with the minor vein boxed in A. Maize veins display Kranz anatomy, with mesophyll cells surrounding bundle sheath cells, which surround the vein. Individual cell types are color coded for identification. Thick-walled sieve elements are shown in pink. C, Suc moves symplastically from mesophyll to bundle sheath to vascular parenchyma cells through plasmodesmata. Suc is exported to the apoplast by vascular parenchyma cells and imported into the companion cell and/or thin-walled sieve element by SUTs (yellow circles). BS, Bundle sheath cells; CC, companion cell; E, epidermal cells; HS, hypodermal sclerenchyma cells; M, mesophyll cells; MX, metaxylem vessels; Ph, phloem cells; PX, protoxylem lacuna; SE, thin-walled sieve element; VP, vascular parenchyma cell; X, xylem tracheary element.
SUC ENTRY INTO A VEIN
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED The majority of grasses are thought to utilize apoplastic phloem loading, based on transmission electron microscopy studies showing an almost complete symplastic isolation of the companion cell-sieve element complex from surrounding cells (Gamalei, 1989). Examples include maize (Evert et al., 1978), barley (Hordeum vulgare; Evert et al., 1996b), and sugarcane (Saccharum hybrid; Robinson-Beers and Evert, 1991). The proposed path of assimilate entry into the phloem in maize is illustrated in Figure 1, B and C. After Suc is synthesized in the cytoplasm of mesophyll cells (Lunn and Furbank, 1999), it diffuses through plasmodesmata into bundle sheath cells and then into vascular parenchyma cells. By an unknown mechanism, the vascular parenchyma cells export Suc to the apoplast. Suc is subsequently imported across the plasma membrane of phloem companion cells and/or sieve elements and then transported by bulk flow to sink tissues. Utilization of an apoplastic phloem-loading mechanism is proposed for C3 grasses, such as barley and wheat (Triticum aestivum; Thompson and Dale, 1981), as well as for the C4 grasses sugarcane and maize. However, the apoplastic loading path may not be universal in grasses, as it has been suggested that rice (Oryza sativa; a C3 species; Kaneko et al., 1980) and Themeda triandra (a C4 species; Botha and Evert, 1988) may instead use symplastic phloem loading, based on the high frequency of plasmodesmata connecting vascular parenchyma cells to companion cells.
Evidence supporting the vascular parenchyma cell as the site of Suc export to the apoplast comes from analysis of the sucrose export defective1 (sxd1) mutant of maize (Russin et al., 1996; Provencher et al., 2001). sxd1 mutants show reduced Suc export capacity and accumulate large quantities of carbohydrates in the distal regions of leaf blades. An examination of the cellular interfaces in these distal regions revealed that callose is ectopically deposited over the cell wall between the bundle sheath and vascular parenchyma cells in leaf minor veins, thereby occluding the plasmodesmata (Botha et al., 2000). This defect is proposed to prevent Suc movement into the vascular parenchyma cells and result in the accumulation of carbohydrates in photosynthetic cells. Sxd1 encodes tocopherol cyclase, the penultimate enzyme in tocopherol (vitamin E) synthesis (Sattler et al., 2003). It is not understood how a lack of tocopherol leads to ectopic callose deposition. However, this function is conserved in potato (Solanum tuberosum) and Arabidopsis (Arabidopsis thaliana), as plants deficient for the Sxd1 orthologous gene show a similar callose-deposition and carbohydrate-accumulation phenotype (Hofius et al., 2004; Maeda et al., 2006).
SUTs
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED Due to the nearly complete symplastic isolation of the sieve element-companion cell complex in the leaf veins of many grasses, Suc entry into the phloem is assumed to require an apoplastic step. SUTs are proposed to mediate Suc transport across the phloem cell plasma membrane; however, until recently, a role for SUTs in phloem loading had not been functionally demonstrated in grasses (see below). SUTs function as Suc-proton symporters with a 1:1 stoichiometry (Bush, 1990; Boorer et al., 1996; Zhou et al., 1997; Carpaneto et al., 2005). The energy to transport Suc against its concentration gradient into the phloem is derived from the proton motive force generated by H+-ATPases in the plasma membrane of phloem cells (Bush, 1993; DeWitt and Sussman, 1995). SUTs contain 12 transmembrane domains that form a pore permitting Suc transport through the membrane. Biochemical studies have shown Suc transport activity for numerous dicot as well as monocot SUTs (for recent reviews, see Aoki et al., 2003; Kühn, 2003; Lalonde et al., 2004; Sauer, 2007). In multiple plants, it has been shown that SUT RNA and/or protein abundance is regulated by Suc (Chiou and Bush, 1998; Aoki et al., 1999; Matsukura et al., 2000; Vaughn et al., 2002; Ransom-Hodgkins et al., 2003), but the genes that control SUT expression have not been identified.
Historically, SUT genes were named in the order in which they were identified; hence, SUT1 in one species was orthologous to SUT5 in another. Furthermore, in Arabidopsis and several other plants, some SUT genes are named SUC for Suc carriers. To clarify their evolutionary relatedness, a phylogeny of different SUT proteins is presented (Fig. 2 ). For simplicity and consistency, we have renamed three grass SUTs according to their orthology with rice SUT genes (Aoki et al., 2003; Fig. 2; Table I ). We propose that all grass SUT genes that are identified in the future be named according to their phylogenetic relationships to avoid confusion.
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Figure 2. Phylogenetic tree of all grass and select dicot SUTs showing relationships among SUTs. The phylogenetic tree was created with PHYLIP based on the alignment of deduced amino acid sequences using the ClustalW program. The tree was rooted with the SUT-like sequence from Aspergillus fumigatus (AfSUT; accession no. EAL92728) as an outgroup. The five different groups of SUTs are indicated with brackets, and the locations of the rice SUTs are highlighted. GenBank accession numbers for grass SUTs are presented in Table I, and those from dicots are as follows: AtSUC1 (At1g71880), AtSUC2 (At1g22710), AtSUC3 (At2g02860), AtSUT4 (At1g09960), AtSUC5 (At1g71890), AtSUC9 (At5g06170), LeSUT1 (X82275), LeSUT2 (AAG12987), LjSUT4 (CAD61275), NtSUT1 (X82276), PmSUC1 (Plantago major; CAI59556), PmSUC2 (X75764), PmSUC3 (CAD58887), PsSUF4 (DQ221697), StSUT1 (CAA48915), and StSUT4 (AAG25923). [See online article for color version of this figure.]
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Table I. Grass SUTs
Based on sequence homology and biochemical activity, SUTs were previously divided into three types: type I was composed exclusively of dicot sequences (e.g. AtSUC2), but type II (OsSUT1 and AtSUC3, which is identical to AtSUT2) and type III (HvSUT2 and AtSUT4, which is identical to AtSUC4) contained both monocot and dicot proteins (Aoki et al., 2003; Lalonde et al., 2004). With increasing numbers of SUT sequences available, phylogenetic analysis divided the SUTs into four clades, splitting the type II subfamily into two groups (Sauer, 2007). The recent completion of the draft genome sequences for sorghum (Sorghum bicolor), Brachypodium distachyon, and maize facilitated the identification of their corresponding SUT sequences, and an additional fifth group emerged (Fig. 2). Groups 1 and 5 (formerly type II) consist entirely of monocot SUTs, group 2 (formerly type I) contains only dicot SUTs, and both group 3 (formerly type II) and group 4 (formerly type III) contain monocot and dicot SUTs (Fig. 2).
The Arabidopsis genome contains the largest SUT gene family characterized to date, with nine SUT-like genes, although two are categorized as pseudogenes (Sauer et al., 2004). Of the remaining seven, five are group 2 members, one is a group 3 member, and the last is a group 4 member. The rice genome contains five SUT genes: two group 1 members and a single gene each from groups 3, 4, and 5 (Aoki et al., 2003; Fig. 2; Table I). Analyses of the draft genomes of maize, sorghum, and Brachypodium indicate that they contain the same five SUT genes as rice (Fig. 2; Table I). In addition, SUT5 appears to have been duplicated in maize and sorghum. As has been observed previously in dicots, in all the grass genomes available, there is only a single group 3 or 4 SUT gene (the two group 3 rice genes reported by Sauer [2007] are alleles from different cultivars). In dicots, the group 2 SUT genes underwent duplication and neofunctionalization (Baud et al., 2005; Sivitz et al., 2007, 2008), and in monocots, the group 1 SUTs similarly expanded (Fig. 2). Furthermore, it is possible that the grass SUT3 sequences, which are well resolved from the SUT1 sequences within group 1, are functionally distinct and may eventually be split into a group 6.
It is important to point out that all data currently available on monocot SUTs are derived solely from grasses. In comparison with dicots, few monocots have been characterized for leaf anatomy and phloem-loading mechanism (Gamalei, 1989). Additionally, more monocot genomes need to be sequenced to better understand the evolution of the SUT gene family and whether SUT functions are conserved in nongrass monocots. Nevertheless, the functions of almost all grass SUT genes still remain to be determined.
FUNCTION OF GROUP 2 SUTS
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED Group 2 SUT family members are unique to dicots. In Arabidopsis, the best characterized group 2 SUT shown to function in phloem loading is AtSUC2. AtSUC2 RNA and protein are expressed in the companion cells of minor veins in a pattern reflective of the sink-to-source transition in leaves (Truernit and Sauer, 1995; Stadler and Sauer, 1996; Wright et al., 2003). AtSUC2 has a biochemical affinity for Suc consistent with a role in phloem loading from the apoplast (Chandran et al., 2003). Definitive proof of a function in phloem loading came from analyses of T-DNA insertion mutations in AtSUC2 (Gottwald et al., 2000; Srivastava et al., 2008). Mutant plants have reduced Suc export from leaves, chlorotic leaves that accumulate carbohydrates, diminished shoot growth, and delayed flowering. Recently, using a tissue-specific promoter to complement an Atsuc2 mutant, it was shown that the only essential function in photoassimilate distribution for AtSUC2 was to load Suc into the phloem in the leaf minor veins (Srivastava et al., 2008).
Phenotypes similar to those displayed by Atsuc2 mutant plants have been reported in transgenic plants expressing antisense RNAs for SUT1 in potato (Riesmeier et al., 1994), tobacco (Nicotiana tabacum; Bürkle et al., 1998), and tomato (Solanum lycopersicum [formerly Lycopersicon esculentum; Le]; Hackel et al., 2006), demonstrating that SUT1 functions in phloem loading in these plants. Solanaceous SUT1 protein was first reported as localizing to the plasma membrane of sieve elements (Kühn et al., 1997; Barker et al., 2000). However, a recent analysis found that SUT1 in tobacco, potato, and tomato localized to the plasma membrane of companion cells, suggesting that both solanaceous and Arabidopsis plants load Suc into the phloem companion cells (Schmitt et al., 2008). Contrary to this, another recent publication reports SUT1 in potato localizing to the sieve elements in stems and leaf minor veins (Krügel et al., 2008). In all of these papers, SUT1 localization was determined by immunofluorescence, which is dependent on the specificities of the different antibodies used and may explain the disparate results (for discussion, see Schmitt et al., 2008). It may be necessary to utilize a different approach to resolve which cells express SUT1 and are responsible for phloem loading in the Solanaceae.
GROUP 3, 4, AND 5 SUT FAMILY MEMBERS
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED The functions of group 3 SUTs are not clear. Based on sequence characteristics and an initial report of a lack of transport activity, it was postulated that AtSUT2/AtSUC3 might function as a Suc sensor (Barker et al., 2000). However, subsequent studies have questioned this hypothesis by finding that null mutations in AtSUC3 have no obvious phenotype and that AtSUC3 protein has measurable biochemical activity (Meyer et al., 2000; Barth et al., 2003). Interestingly, the gene is expressed in many sink tissues and is strongly induced by wounding (Meyer et al., 2000, 2004). The only report characterizing a group 3 SUT in grasses showed that the rice OsSUT4 gene is expressed in all tissues examined, although at higher levels in sink leaves (Aoki et al., 2003). More research is needed to understand the function of these SUTs in both grasses and dicot plants.
The first member of the group 4 SUT subfamily identified was HvSUT2 (Weschke et al., 2000). HvSUT2 has Suc transport activity when expressed in yeast. The gene is expressed at the highest level in developing leaves and at approximately equal levels in roots, mature leaves, developing grains, anthers, and gynoecium tissues. Based on RNA expression, it was proposed that HvSUT2 may play a general housekeeping role.
Endler et al. (2006) discovered that HvSUT2 was a vacuolar membrane-resident protein through a proteomic analysis of tonoplasts isolated from barley leaf mesophyll cells. Similar analyses showed that AtSUT4, the Arabidopsis homolog, was also a tonoplast-localized protein. These data were confirmed by HvSUT2-GFP and AtSUT4-GFP transient subcellular localization studies in onion (Allium cepa) bulb and Arabidopsis leaf epidermal cells. Based on their localization, it was suggested that these proteins function in Suc exchange between the vacuole and cytoplasm. Hence, the Suc uptake activity measured in yeast for these SUTs was likely due to mistargeting to the plasma membrane (Weise et al., 2000; Weschke et al., 2000).
Recently, Reinders et al. (2008) found that the Lotus japonicus group 4 homolog, LjSUT4, also localized to the vacuolar membrane, indicating that LjSUT4 may likewise function to transport Suc from the vacuole into the cytoplasm. Using Xenopus laevis oocytes, it was determined that LjSUT4 was a proton-coupled low-affinity Suc transporter. As the authors point out, fortuitous mislocalization of the tonoplast-resident protein to the oocyte plasma membrane permitted electrophysiological analyses. As oocytes do not contain vacuoles, it remains a possibility that the biochemical properties determined for the transporter do not completely reflect its endogenous role. It will be interesting to determine the function of the protein within its native context.
The SUT homolog PsSUF4 from pea (Pisum sativum), showing 73% amino acid identity to LjSUT4, was characterized by heterologous expression in yeast and shown to be a Suc facilitator, rather than a symporter, that functions independently of a H+ gradient (Zhou et al., 2007). The subcellular localization of PsSUF4 in plants was not reported, but if similarly localized to the tonoplast, it may be responsible for Suc entry into the vacuole. If so, this suggests the intriguing possibility that PsSUF4 (and maybe other group 4 SUTs) may facilitate Suc transport into and out of vacuoles. In agreement with this hypothesis, Suc uptake studies in barley leaf mesophyll cell vacuoles found that Suc uptake was not energy dependent and that transport was not influenced by protonophores, which destroy the H+ gradient across the membrane (Kaiser and Heber, 1984). These data suggest that vacuolar import of Suc occurred by facilitated diffusion rather than active transport. Clearly, more work is needed to test the functions of these genes in plants to understand their role in transitory Suc storage in vacuoles.
The sole report on the in vivo function of a group 4 SUT concerns StSUT4 from potato (Chincinska et al., 2008). StSUT4 localized to both the plasma membrane and the endomembranes surrounding the nucleus in tobacco and potato leaves, but not to the tonoplast. This suggests that not all group 4 SUTs function in the tonoplast and that another SUT in potato may function to transport Suc between the vacuole and cytoplasm. Antisense reduction of StSUT4 expression led to altered accumulation of soluble sugars in leaves and increased phloem export of Suc. These phenotypes are the opposite of those observed in antisense StSUT1 plants (Bürkle et al., 1998), suggesting that StSUT4 has a distinct biological function. Additionally, antisense StSUT4 plants flowered early, had higher tuber production, and had reduced shade avoidance. The authors proposed that StSUT4 may act in the interplay of carbon availability and flower induction pathways.
Very little is known about other grass members of the group 4 SUTs. The rice ortholog of HvSUT2 is OsSUT2. From reverse transcription-PCR experiments, OsSUT2 is constitutively expressed in vegetative and reproductive tissues, although expression decreases toward the end of seed development (Aoki et al., 2003). Determining the physiological functions of these genes awaits the characterization of plants containing the respective loss-of-function mutations.
Similarly, the functions of group 5 SUTs are unknown and remain to be determined. OsSUT5 is expressed nearly ubiquitously and shows the highest expression level in sink leaves (Aoki et al., 2003).
FUNCTIONS OF GROUP 1 SUTs
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED Group 1 SUTs are present only in monocots and are subdivided into two clades represented by OsSUT1 and OsSUT3 (Fig. 2). Nothing is currently known of the functions of grass SUT3 genes. Here, we briefly summarize what is known about the functions of grass SUT1 genes in phloem loading in leaves as well as their roles in other tissues.
Rice
OsSUT1 was the first SUT cloned from monocots (Hirose et al., 1997). Reverse transcription-PCR shows that it is expressed at high levels in germinating seeds, source leaf sheaths, panicles, and developing grains and at lower levels in sink and source leaves (Aoki et al., 2003). OsSUT1 transcript was expressed in companion cells in leaf sheaths and in the scutellar vasculature of germinating seeds (Matsukura et al., 2000; Furbank et al., 2001; Scofield et al., 2007a). OsSUT1 may also play a role in maternal tissues, as both transcript and protein have been localized to the nucellus, vascular parenchyma tissue, and nucellar projection (Furbank et al., 2001). Promoter:GUS analyses and immunolocalization experiments showed that OsSUT1 was expressed in the companion cells and sieve elements, where it may function in phloem loading of Suc for transport to seedling shoots and roots (Scofield et al., 2007a). OsSUT1 is also expressed in the phloem along the long-distance transport path from the flag leaf blade to the base of the filling grain (Scofield et al., 2007b). It was suggested that OsSUT1 may function in Suc retrieval from the apoplasm along the transport path.
OsSUT1 does not appear to have an essential function in phloem loading of Suc in mature leaf blades. This was shown by strongly reducing OsSUT1 gene expression by antisense RNA suppression (Ishimaru et al., 2001; Scofield et al., 2002). Antisense lines with almost complete reduction in RNA or protein levels had normal vegetative growth and no alteration in photosynthesis or leaf carbohydrate contents, contrary to what would be expected if OsSUT1 principally functioned in phloem loading. Instead, transgenic plants had reduced grain filling and decreased germination, indicating that OsSUT1 plays an important role in transporting Suc to the developing grain and in remobilizing stored carbohydrates during early seedling growth. No visible phenotype was observed in leaves of OsSUT1 antisense lines, potentially due to genetic redundancy (Ishimaru et al., 2001; Aoki et al., 2003) or to utilization of a symplastic phloem-loading pathway (Kaneko et al., 1980). To our knowledge, these two reports are the only publications to date that directly assess the biological function of a grass SUT in vivo.
In support of a symplastic loading pathway in rice, fluorescent dyes were recently used to show that a xylem sap retrieval pathway functions in rice leaf blades to transfer solutes from the xylem transpiration stream into adjacent vascular parenchyma cells and into the phloem sieve elements (Botha et al., 2008). Potentially related to a functional symplastic phloem-loading pathway, grass veins contain two types of sieve elements: thick walled and thin walled (Kuo and O'Brien, 1974; Walsh, 1974; Evert et al., 1978; Botha, 2005). Botha and coworkers (2008) showed that the thick-walled sieve elements were connected by plasmodesmata to vascular parenchyma cells and accumulated significant amounts of fluorescent dye. In contrast, the thin-walled sieve elements and their companion cells were virtually symplastically isolated from other cells and contained much less dye. An intriguing possibility that needs more attention is that grass leaves may be able to use both symplastic (via thick-walled sieve elements?) and apoplastic (thin-walled sieve elements) phloem-loading mechanisms (Chonan et al., 1985; van Bel, 1993; Botha, 2005). Perhaps if the apoplastic pathway is not functional, the ability to symplastically load Suc into the phloem sustains plant growth. In fact, Srivastava et al. (2008) recently discussed the possibility that Arabidopsis may similarly be able to load Suc directly into the phloem using a completely symplastic pathway. Hence, it is possible that plants thought to use apoplastic phloem loading based on anatomical considerations are also conditional symplastic loaders (for discussion of mixed loading, see van Bel, 1993).
Wheat
In wheat, three homeologous genes known as TaSUT1A, TaSUT1B, and TaSUT1D (corresponding to the A, B, and D progenitor genomes that make up the genome of hexaploid wheat) have been characterized (Aoki et al., 2002). The three are almost equally expressed in leaves, internodes, and developing grains (Aoki et al., 2004). RNA in situ hybridization showed that the TaSUT1 RNA accumulated specifically in companion cells in leaves. However, immunolocalization experiments with an antibody raised against the rice OsSUT1 protein determined that TaSUT1 is localized to the plasma membrane of sieve elements in both leaves and stem tissues. This suggests that phloem loading of Suc occurs in the sieve elements in wheat and that the protein trafficks through plasmodesmata (Aoki et al., 2004). Based on its expression pattern, TaSUT1 has been proposed to function in phloem loading in source leaves and in retrieval of leaked Suc along the transport phloem in sheath and stem tissues. In developing grains, TaSUT1 localized to the plasma membrane of modified aleurone and subaleurone cells adjacent to the endosperm cavity, where it has been proposed to play a role in Suc uptake into filial tissues (Bagnall et al., 2000; Aoki et al., 2006).
Barley
HvSUT1 in barley is expressed at very high levels in developing seeds during the time of starch deposition (Weschke et al., 2000). RNA in situ hybridization determined that HvSUT1 transcript mainly accumulated in the maternal nucellar projection and in the endosperm transfer layer, which correspond to the site of Suc exchange between maternal and filial tissues. HvSUT1 is proposed to function in developing seeds in Suc uptake for delivery to starchy endosperm cells and in Suc retrieval in maternal tissue. HvSUT1 is expressed at lower levels in sink and source leaves, but its function in these tissues is not known. HvSUT1 RNA has been detected in phloem sap, indicating that the RNA trafficked through plasmodesmata from the companion cells to sieve elements (Doering-Saad et al., 2002). Expression of HvSUT1 in Xenopus oocytes determined that the transporter had moderate Suc affinity and was more substrate selective than AtSUC2 (Sivitz et al., 2005).
Sugarcane
Sugarcane internodes have the remarkable ability to store massive amounts of Suc, depositing it in both the vacuoles of stem storage parenchyma cells and the apoplast surrounding these cells (Welbaum and Meinzer, 1990). To prevent apoplastic backflow of Suc to the phloem, the stem veins are surrounded by sclerenchyma cells, which contain suberin and lignin in their cell walls (Walsh et al., 2005). This produces an apoplastic barrier to solute movement, as shown by the fact that xylem sap contains no Suc (Welbaum et al., 1992). Therefore, Suc must take an entirely symplastic route from the phloem into the storage parenchyma cells, where it is exported to the apoplast. Fluorescent dye tracer studies support this transport path (Rae et al., 2005; Walsh et al., 2005).
ShSUT1 RNA is expressed at the highest levels in tissues experiencing high sugar flux, specifically, mature exporting leaves and sugar-accumulating internodes (Rae et al., 2005). Immunolocalization experiments with an antibody raised against a ShSUT1 peptide indicated that the protein is not expressed in the phloem and therefore does not function in phloem loading or retrieval in these tissues (Rae et al., 2005). Instead, ShSUT1 localized to vascular parenchyma and mestome sheath cells (inner bundle sheath cell layer) in leaf and stem tissues. It was proposed that ShSUT1 may function as a biochemical barrier to apoplastic transport of Suc and retrieve any Suc leaked to the apoplast (Rae et al., 2005). Consistent with this idea, electrophysiological studies of ShSUT1 in oocytes determined that ShSUT1 had the highest selectivity for Suc of any SUT known, along with a relatively low Suc affinity (Reinders et al., 2006). It is likely that additional SUTs function in phloem loading of Suc in sugarcane, as an antibody raised against rice OsSUT1 immunolocalized to phloem tissue in leaves and stem (Rae et al., 2005). Further studies are needed to characterize additional sugarcane SUTs to determine which family member(s) function in the phloem.
Maize
ZmSUT1 is highly expressed in photosynthetic tissues, with maximal expression in leaf blades at the end of the day and minimal expression during the night (Aoki et al., 1999). Additionally, in an expanding leaf, ZmSUT1 is expressed in a gradient, with highest levels in mature tissues at the tip and lowest levels in the immature base, a pattern reflective of the sink-to-source transition. Based on its expression pattern and biochemical activity, it has been proposed that ZmSUT1 functions in phloem loading in source tissues (Aoki et al., 1999; Carpaneto et al., 2005). However, as the closely related genes OsSUT1 and ShSUT1 (Fig. 2) apparently do not function in phloem loading, it is not certain what role ZmSUT1 plays in planta. Complicating the matter, ZmSUT1 has also been shown to be capable of transporting Suc across the plasma membrane in both directions in Xenopus oocytes if the membrane potential, Suc, and pH gradients are reversed (Carpaneto et al., 2005).
We recently determined that ZmSUT1 functions in phloem loading by characterizing a knockout mutation (Slewinski et al., 2009). Zmsut1 mutant plants developed chlorotic leaves that hyperaccumulated carbohydrates and prematurely senesced. Application of [14C]Suc to abraded leaves demonstrated that Suc export was greatly diminished in Zmsut1 mutants compared to wild type. In addition, mutant plants had reduced plant height, fewer leaves, delayed flowering, and stunted tassel development. Presumably, these phenotypes result from a reduction in assimilates delivered to sink tissues due to the failure to export Suc from source leaves. These phenotypes are similar to those reported in dicot plants containing mutations in SUT genes that are responsible for phloem loading (Riesmeier et al., 1994; Bürkle et al., 1998; Gottwald et al., 2000; Hackel et al., 2006; Srivastava et al., 2008). Hence, it appears, at least in maize, that SUT1 function is essential for phloem loading of Suc. Determining the biological functions of the five additional SUTs in maize (Fig. 2) will require characterizing plants that harbor mutations in each gene.
Tie-dyed Loci
In addition to Sxd1 and ZmSUT1, several other maize genes that affect carbohydrate accumulation in leaves have been characterized. The tie-dyed1 (tdy1) mutant was identified by its variegated yellow and green leaf phenotype (Braun et al., 2006). tdy1 sectors violate the clonal cell lineages in maize leaves, suggesting that a mobile signal is responsible for sector formation. Phenotypic characterization of the mutant revealed that the tdy1 yellow sectors hyperaccumulated soluble sugars and starch, indicating that carbon partitioning was perturbed. Additionally, tdy1 mutants showed reduced plant stature, delayed flowering, and decreased yield, phenotypes that resemble dicot and maize SUT mutants (see above). However, because tdy1 mutants are variegated, it was proposed that the gene functions as an osmotic stress or sugar sensor to promote phloem loading, rather than acting directly to transport sugars (Braun et al., 2006). Through a clonal mosaic analysis, Tdy1 function was mapped to the middle layer of leaves, consisting of the interveinal mesophyll, bundle sheath, and vascular cells (Baker and Braun, 2007). To further dissect the function of Tdy1 in carbohydrate partitioning, epistasis analysis was used to determine that Tdy1 does not play a role in the photosynthetic cells in starch metabolism (Slewinski et al., 2008). In addition, it was shown that Tdy1 functions independently of Sxd1 in controlling carbohydrate accumulation in leaves (Ma et al., 2008). However, based on its nearly identical mutant phenotype and a dosage-sensitive genetic interaction, Tdy1 was found to function in the same pathway as Tdy2 (Baker and Braun, 2008). It was hypothesized that TDY1 and TDY2 may form a protein complex that promotes phloem loading.
To understand the function of Tdy1, we recently cloned the gene (Ma et al., 2009). Tdy1 encodes a novel transmembrane protein found only in grasses, although two conserved protein subdomains are present in monocots and dicots. RNA in situ hybridization studies determined that Tdy1 RNA was expressed exclusively in phloem cells. In fact, Tdy1 is expressed as soon as protophloem cells mature, indicating that Tdy1 may be a useful marker for early phloem cell differentiation. Monitoring symplastic solute movement with a fluorescent dye in wild-type and mutant leaves revealed that phloem loading was impaired in tdy1 mutants. Therefore, it was proposed that Tdy1 may function to promote phloem loading, potentially through modulating the activity of a SUT. Future investigations to examine the genetic and biochemical interactions between TDY1 and maize SUTs will test this hypothesis.
CONCLUDING THOUGHTS
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED Our understanding of the genetic regulation of carbon partitioning is just beginning. In addition to the genes described above, we have identified many other maize loci that regulate leaf carbohydrate accumulation. Characterization of the function of these genes and all SUT family members will open up new avenues of investigation into the control of carbon partitioning in plants. Moreover, it will afford exciting prospects for biotechnological approaches to enhance crop yield and biofuels production. As a case in point, it was recently demonstrated that the sugar content of sugarcane stems could be doubled with no apparent defect to plant growth, illustrating the potential to greatly modify carbon partitioning patterns (Wu and Birch, 2007). In addition, hyperaccumulation of carbohydrates in maize leaves redirected significant amounts of carbon to cellulose in the cell wall (Baker and Braun, 2008). These examples indicate that tremendous opportunities exist to manipulate carbon partitioning pathways in grasses. Additional research is needed to uncover the genetic pathways and control points regulating carbohydrate partitioning in plants to realize these goals.
ACKNOWLEDGMENTS We thank John Ward, two anonymous reviewers, and members of the Braun and McSteen laboratories for valuable comments that improved the manuscript. We gratefully acknowledge the Maize Genome Sequencing project and the Sorghum and Brachypodium Genome Projects of the Department of Energy Joint Genome Institute for making the draft genome sequences available prior to publication. We sincerely appreciate the efforts of Toby Kellogg and Robin Buell to organize this Focus Issue on the Grasses.
Received September 1, 2008; accepted October 19, 2008; published January 7, 2009.
FOOTNOTES 1 This work was supported by the National Research Initiative of the U.S. Department of Agriculture Cooperative State Research, Education, and Extension Service (grant no. 2008–35304–04597 to D.M.B.).
The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: David M. Braun (dbraun@psu.edu
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[C] Some figures in this article are displayed in color online but in black and white in the print edition.
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* Corresponding author; e-mail dbraun@psu.edu
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Plants have specialized organs for distinct functions. Leaves perform photosynthesis and fix carbon, whereas roots absorb water and minerals. To distribute resources between these organs, plants have a vasculature composed of phloem and xylem. The xylem conducts water and minerals from the roots up to the shoots. The phloem transports carbon- and nitrogen-containing compounds from mature leaves to the roots and to other nonphotosynthetic organs such as flowers and fruits. Phloem tissue comprises two main cell types: sieve elements and companion cells. Sieve elements conduct nutrients, while companion cells metabolically support the sieve elements (van Bel and Knoblauch, 2000). The vascular system represents a highly integrated distribution network essential not only for the life of the plant but also for the life of the planet, as nearly all terrestrially produced chemical energy, including our food supply, is derived from plants.
Photosynthesis and carbon assimilation occur in leaf mesophyll cells and additionally in bundle sheath cells in C4 plants. For distribution to distal tissues, fixed carbon must move out of the photosynthetic cells and into the phloem. If the photoassimilates (assimilated carbon) diffuse down a concentration gradient from the mesophyll cells into the phloem following an entirely cytoplasmic path through plasmodesmata (intercellular channels through which small molecules freely diffuse), it is referred to as symplastic phloem loading (Turgeon and Medville, 1998). However, if the assimilated carbon must cross a membrane prior to entering into the phloem, it is called apoplastic phloem loading (van Bel, 1993; Turgeon, 2006). In this case, the concentration of the transported assimilate is higher in the phloem than in the photosynthetic cells. Because apoplastic phloem loading involves movement against a concentration gradient, it requires energy in the form of a pH gradient generated by H+-ATPases (Bush, 1993; Gaxiola et al., 2007).
Carbon partitioning is the process whereby assimilates are distributed throughout the plant body from photosynthetic tissues. For most plants, this occurs by loading Suc into the phloem and transporting it from source tissues (net exporters) to sink tissues (net importers), where Suc is unloaded (Turgeon, 1989; van Bel, 2003). This process is well characterized at the physiological, biochemical, and anatomical levels (Hofstra and Nelson, 1969; Fellows and Geiger, 1974; Evert et al., 1978; Nguyen-Quoc et al., 1990; Huber and Hanson, 1992; Evert et al., 1996a; Koch, 1996; Paul and Foyer, 2001). However, despite the obvious importance of this process for plant growth and development, few genes that function in carbon partitioning have been identified. Suc, K+, and water transporters/channels have been characterized for their contribution to the transport of Suc in the phloem (Deeken et al., 2002; Lalonde et al., 2004; Maurel, 2007; Sauer, 2007). Of these, the best described genes that directly control Suc loading into the phloem encode Suc transporters (SUTs; Lalonde et al., 2004; Sauer, 2007).
In this review, we discuss phloem loading and the control of carbon partitioning in grasses, focusing on SUTs and highlighting similarities and differences with eudicots. Additionally, we cover related aspects of phloem loading, such as leaf anatomy, and discuss other genes regulating carbohydrate accumulation in grass leaves. For additional discussions of genes that function in phloem loading and carbon partitioning (e.g. H+-ATPase, K+ channel, Suc synthase, aquaporins), we refer interested readers to the following articles (Nolte and Koch, 1993; DeWitt and Sussman, 1995; Hannah et al., 2001; Deeken et al., 2002; Dinges et al., 2003; Ma et al., 2004; Hardin et al., 2006; Lu and Sharkey, 2006; Smith and Stitt, 2007).
LEAF ANATOMY
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED As the path of phloem loading is intimately related to leaf structure, we begin with a brief overview of grass leaf anatomy, describing a maize (Zea mays) leaf blade as a typical example (Fig. 1A ). Three orders of veins are arranged longitudinally along the main axis of the leaf (Esau, 1977). The largest type contains large metaxylem vessels and a protoxylem lacuna (space; Fig. 1A). Additionally, as structural support, hypodermal sclerenchyma girders are located between the vein and the epidermal cell layers. Intermediate veins lack metaxylem vessels but have hypodermal sclerenchyma caps on one or both sides of the vein. The smallest veins have neither metaxylem vessels nor hypodermal sclerenchyma caps. Collectively, the intermediate and small veins are referred to as minor veins, while the largest veins are known as lateral or major veins. Minor veins intergrade into the lateral veins (Russell and Evert, 1985). Transverse veins with anatomy similar to the small veins connect adjacent longitudinal veins. All veins contain phloem cells that transport photoassimilates, but the various orders of veins play different roles in carbon partitioning. Minor veins are the site where photoassimilates are loaded into the vein (Fritz et al., 1983). Lateral veins principally function in the long-distance transport of assimilated carbon out of the leaf into the stem (Fritz et al., 1989). Transverse veins shunt assimilates from minor veins to lateral veins (Fritz et al., 1989).
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Figure 1. Maize leaf anatomy and path of Suc movement. A, Cross section of a maize leaf blade viewed under UV light to visualize cellular anatomy. Chlorophyll autofluoresces red and cell walls autofluoresce blue. One lateral vein and three minor veins are shown. B, Schematic diagram of cell types associated with the minor vein boxed in A. Maize veins display Kranz anatomy, with mesophyll cells surrounding bundle sheath cells, which surround the vein. Individual cell types are color coded for identification. Thick-walled sieve elements are shown in pink. C, Suc moves symplastically from mesophyll to bundle sheath to vascular parenchyma cells through plasmodesmata. Suc is exported to the apoplast by vascular parenchyma cells and imported into the companion cell and/or thin-walled sieve element by SUTs (yellow circles). BS, Bundle sheath cells; CC, companion cell; E, epidermal cells; HS, hypodermal sclerenchyma cells; M, mesophyll cells; MX, metaxylem vessels; Ph, phloem cells; PX, protoxylem lacuna; SE, thin-walled sieve element; VP, vascular parenchyma cell; X, xylem tracheary element.
SUC ENTRY INTO A VEIN
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED The majority of grasses are thought to utilize apoplastic phloem loading, based on transmission electron microscopy studies showing an almost complete symplastic isolation of the companion cell-sieve element complex from surrounding cells (Gamalei, 1989). Examples include maize (Evert et al., 1978), barley (Hordeum vulgare; Evert et al., 1996b), and sugarcane (Saccharum hybrid; Robinson-Beers and Evert, 1991). The proposed path of assimilate entry into the phloem in maize is illustrated in Figure 1, B and C. After Suc is synthesized in the cytoplasm of mesophyll cells (Lunn and Furbank, 1999), it diffuses through plasmodesmata into bundle sheath cells and then into vascular parenchyma cells. By an unknown mechanism, the vascular parenchyma cells export Suc to the apoplast. Suc is subsequently imported across the plasma membrane of phloem companion cells and/or sieve elements and then transported by bulk flow to sink tissues. Utilization of an apoplastic phloem-loading mechanism is proposed for C3 grasses, such as barley and wheat (Triticum aestivum; Thompson and Dale, 1981), as well as for the C4 grasses sugarcane and maize. However, the apoplastic loading path may not be universal in grasses, as it has been suggested that rice (Oryza sativa; a C3 species; Kaneko et al., 1980) and Themeda triandra (a C4 species; Botha and Evert, 1988) may instead use symplastic phloem loading, based on the high frequency of plasmodesmata connecting vascular parenchyma cells to companion cells.
Evidence supporting the vascular parenchyma cell as the site of Suc export to the apoplast comes from analysis of the sucrose export defective1 (sxd1) mutant of maize (Russin et al., 1996; Provencher et al., 2001). sxd1 mutants show reduced Suc export capacity and accumulate large quantities of carbohydrates in the distal regions of leaf blades. An examination of the cellular interfaces in these distal regions revealed that callose is ectopically deposited over the cell wall between the bundle sheath and vascular parenchyma cells in leaf minor veins, thereby occluding the plasmodesmata (Botha et al., 2000). This defect is proposed to prevent Suc movement into the vascular parenchyma cells and result in the accumulation of carbohydrates in photosynthetic cells. Sxd1 encodes tocopherol cyclase, the penultimate enzyme in tocopherol (vitamin E) synthesis (Sattler et al., 2003). It is not understood how a lack of tocopherol leads to ectopic callose deposition. However, this function is conserved in potato (Solanum tuberosum) and Arabidopsis (Arabidopsis thaliana), as plants deficient for the Sxd1 orthologous gene show a similar callose-deposition and carbohydrate-accumulation phenotype (Hofius et al., 2004; Maeda et al., 2006).
SUTs
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED Due to the nearly complete symplastic isolation of the sieve element-companion cell complex in the leaf veins of many grasses, Suc entry into the phloem is assumed to require an apoplastic step. SUTs are proposed to mediate Suc transport across the phloem cell plasma membrane; however, until recently, a role for SUTs in phloem loading had not been functionally demonstrated in grasses (see below). SUTs function as Suc-proton symporters with a 1:1 stoichiometry (Bush, 1990; Boorer et al., 1996; Zhou et al., 1997; Carpaneto et al., 2005). The energy to transport Suc against its concentration gradient into the phloem is derived from the proton motive force generated by H+-ATPases in the plasma membrane of phloem cells (Bush, 1993; DeWitt and Sussman, 1995). SUTs contain 12 transmembrane domains that form a pore permitting Suc transport through the membrane. Biochemical studies have shown Suc transport activity for numerous dicot as well as monocot SUTs (for recent reviews, see Aoki et al., 2003; Kühn, 2003; Lalonde et al., 2004; Sauer, 2007). In multiple plants, it has been shown that SUT RNA and/or protein abundance is regulated by Suc (Chiou and Bush, 1998; Aoki et al., 1999; Matsukura et al., 2000; Vaughn et al., 2002; Ransom-Hodgkins et al., 2003), but the genes that control SUT expression have not been identified.
Historically, SUT genes were named in the order in which they were identified; hence, SUT1 in one species was orthologous to SUT5 in another. Furthermore, in Arabidopsis and several other plants, some SUT genes are named SUC for Suc carriers. To clarify their evolutionary relatedness, a phylogeny of different SUT proteins is presented (Fig. 2 ). For simplicity and consistency, we have renamed three grass SUTs according to their orthology with rice SUT genes (Aoki et al., 2003; Fig. 2; Table I ). We propose that all grass SUT genes that are identified in the future be named according to their phylogenetic relationships to avoid confusion.
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Figure 2. Phylogenetic tree of all grass and select dicot SUTs showing relationships among SUTs. The phylogenetic tree was created with PHYLIP based on the alignment of deduced amino acid sequences using the ClustalW program. The tree was rooted with the SUT-like sequence from Aspergillus fumigatus (AfSUT; accession no. EAL92728) as an outgroup. The five different groups of SUTs are indicated with brackets, and the locations of the rice SUTs are highlighted. GenBank accession numbers for grass SUTs are presented in Table I, and those from dicots are as follows: AtSUC1 (At1g71880), AtSUC2 (At1g22710), AtSUC3 (At2g02860), AtSUT4 (At1g09960), AtSUC5 (At1g71890), AtSUC9 (At5g06170), LeSUT1 (X82275), LeSUT2 (AAG12987), LjSUT4 (CAD61275), NtSUT1 (X82276), PmSUC1 (Plantago major; CAI59556), PmSUC2 (X75764), PmSUC3 (CAD58887), PsSUF4 (DQ221697), StSUT1 (CAA48915), and StSUT4 (AAG25923). [See online article for color version of this figure.]
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Table I. Grass SUTs
Based on sequence homology and biochemical activity, SUTs were previously divided into three types: type I was composed exclusively of dicot sequences (e.g. AtSUC2), but type II (OsSUT1 and AtSUC3, which is identical to AtSUT2) and type III (HvSUT2 and AtSUT4, which is identical to AtSUC4) contained both monocot and dicot proteins (Aoki et al., 2003; Lalonde et al., 2004). With increasing numbers of SUT sequences available, phylogenetic analysis divided the SUTs into four clades, splitting the type II subfamily into two groups (Sauer, 2007). The recent completion of the draft genome sequences for sorghum (Sorghum bicolor), Brachypodium distachyon, and maize facilitated the identification of their corresponding SUT sequences, and an additional fifth group emerged (Fig. 2). Groups 1 and 5 (formerly type II) consist entirely of monocot SUTs, group 2 (formerly type I) contains only dicot SUTs, and both group 3 (formerly type II) and group 4 (formerly type III) contain monocot and dicot SUTs (Fig. 2).
The Arabidopsis genome contains the largest SUT gene family characterized to date, with nine SUT-like genes, although two are categorized as pseudogenes (Sauer et al., 2004). Of the remaining seven, five are group 2 members, one is a group 3 member, and the last is a group 4 member. The rice genome contains five SUT genes: two group 1 members and a single gene each from groups 3, 4, and 5 (Aoki et al., 2003; Fig. 2; Table I). Analyses of the draft genomes of maize, sorghum, and Brachypodium indicate that they contain the same five SUT genes as rice (Fig. 2; Table I). In addition, SUT5 appears to have been duplicated in maize and sorghum. As has been observed previously in dicots, in all the grass genomes available, there is only a single group 3 or 4 SUT gene (the two group 3 rice genes reported by Sauer [2007] are alleles from different cultivars). In dicots, the group 2 SUT genes underwent duplication and neofunctionalization (Baud et al., 2005; Sivitz et al., 2007, 2008), and in monocots, the group 1 SUTs similarly expanded (Fig. 2). Furthermore, it is possible that the grass SUT3 sequences, which are well resolved from the SUT1 sequences within group 1, are functionally distinct and may eventually be split into a group 6.
It is important to point out that all data currently available on monocot SUTs are derived solely from grasses. In comparison with dicots, few monocots have been characterized for leaf anatomy and phloem-loading mechanism (Gamalei, 1989). Additionally, more monocot genomes need to be sequenced to better understand the evolution of the SUT gene family and whether SUT functions are conserved in nongrass monocots. Nevertheless, the functions of almost all grass SUT genes still remain to be determined.
FUNCTION OF GROUP 2 SUTS
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED Group 2 SUT family members are unique to dicots. In Arabidopsis, the best characterized group 2 SUT shown to function in phloem loading is AtSUC2. AtSUC2 RNA and protein are expressed in the companion cells of minor veins in a pattern reflective of the sink-to-source transition in leaves (Truernit and Sauer, 1995; Stadler and Sauer, 1996; Wright et al., 2003). AtSUC2 has a biochemical affinity for Suc consistent with a role in phloem loading from the apoplast (Chandran et al., 2003). Definitive proof of a function in phloem loading came from analyses of T-DNA insertion mutations in AtSUC2 (Gottwald et al., 2000; Srivastava et al., 2008). Mutant plants have reduced Suc export from leaves, chlorotic leaves that accumulate carbohydrates, diminished shoot growth, and delayed flowering. Recently, using a tissue-specific promoter to complement an Atsuc2 mutant, it was shown that the only essential function in photoassimilate distribution for AtSUC2 was to load Suc into the phloem in the leaf minor veins (Srivastava et al., 2008).
Phenotypes similar to those displayed by Atsuc2 mutant plants have been reported in transgenic plants expressing antisense RNAs for SUT1 in potato (Riesmeier et al., 1994), tobacco (Nicotiana tabacum; Bürkle et al., 1998), and tomato (Solanum lycopersicum [formerly Lycopersicon esculentum; Le]; Hackel et al., 2006), demonstrating that SUT1 functions in phloem loading in these plants. Solanaceous SUT1 protein was first reported as localizing to the plasma membrane of sieve elements (Kühn et al., 1997; Barker et al., 2000). However, a recent analysis found that SUT1 in tobacco, potato, and tomato localized to the plasma membrane of companion cells, suggesting that both solanaceous and Arabidopsis plants load Suc into the phloem companion cells (Schmitt et al., 2008). Contrary to this, another recent publication reports SUT1 in potato localizing to the sieve elements in stems and leaf minor veins (Krügel et al., 2008). In all of these papers, SUT1 localization was determined by immunofluorescence, which is dependent on the specificities of the different antibodies used and may explain the disparate results (for discussion, see Schmitt et al., 2008). It may be necessary to utilize a different approach to resolve which cells express SUT1 and are responsible for phloem loading in the Solanaceae.
GROUP 3, 4, AND 5 SUT FAMILY MEMBERS
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED The functions of group 3 SUTs are not clear. Based on sequence characteristics and an initial report of a lack of transport activity, it was postulated that AtSUT2/AtSUC3 might function as a Suc sensor (Barker et al., 2000). However, subsequent studies have questioned this hypothesis by finding that null mutations in AtSUC3 have no obvious phenotype and that AtSUC3 protein has measurable biochemical activity (Meyer et al., 2000; Barth et al., 2003). Interestingly, the gene is expressed in many sink tissues and is strongly induced by wounding (Meyer et al., 2000, 2004). The only report characterizing a group 3 SUT in grasses showed that the rice OsSUT4 gene is expressed in all tissues examined, although at higher levels in sink leaves (Aoki et al., 2003). More research is needed to understand the function of these SUTs in both grasses and dicot plants.
The first member of the group 4 SUT subfamily identified was HvSUT2 (Weschke et al., 2000). HvSUT2 has Suc transport activity when expressed in yeast. The gene is expressed at the highest level in developing leaves and at approximately equal levels in roots, mature leaves, developing grains, anthers, and gynoecium tissues. Based on RNA expression, it was proposed that HvSUT2 may play a general housekeeping role.
Endler et al. (2006) discovered that HvSUT2 was a vacuolar membrane-resident protein through a proteomic analysis of tonoplasts isolated from barley leaf mesophyll cells. Similar analyses showed that AtSUT4, the Arabidopsis homolog, was also a tonoplast-localized protein. These data were confirmed by HvSUT2-GFP and AtSUT4-GFP transient subcellular localization studies in onion (Allium cepa) bulb and Arabidopsis leaf epidermal cells. Based on their localization, it was suggested that these proteins function in Suc exchange between the vacuole and cytoplasm. Hence, the Suc uptake activity measured in yeast for these SUTs was likely due to mistargeting to the plasma membrane (Weise et al., 2000; Weschke et al., 2000).
Recently, Reinders et al. (2008) found that the Lotus japonicus group 4 homolog, LjSUT4, also localized to the vacuolar membrane, indicating that LjSUT4 may likewise function to transport Suc from the vacuole into the cytoplasm. Using Xenopus laevis oocytes, it was determined that LjSUT4 was a proton-coupled low-affinity Suc transporter. As the authors point out, fortuitous mislocalization of the tonoplast-resident protein to the oocyte plasma membrane permitted electrophysiological analyses. As oocytes do not contain vacuoles, it remains a possibility that the biochemical properties determined for the transporter do not completely reflect its endogenous role. It will be interesting to determine the function of the protein within its native context.
The SUT homolog PsSUF4 from pea (Pisum sativum), showing 73% amino acid identity to LjSUT4, was characterized by heterologous expression in yeast and shown to be a Suc facilitator, rather than a symporter, that functions independently of a H+ gradient (Zhou et al., 2007). The subcellular localization of PsSUF4 in plants was not reported, but if similarly localized to the tonoplast, it may be responsible for Suc entry into the vacuole. If so, this suggests the intriguing possibility that PsSUF4 (and maybe other group 4 SUTs) may facilitate Suc transport into and out of vacuoles. In agreement with this hypothesis, Suc uptake studies in barley leaf mesophyll cell vacuoles found that Suc uptake was not energy dependent and that transport was not influenced by protonophores, which destroy the H+ gradient across the membrane (Kaiser and Heber, 1984). These data suggest that vacuolar import of Suc occurred by facilitated diffusion rather than active transport. Clearly, more work is needed to test the functions of these genes in plants to understand their role in transitory Suc storage in vacuoles.
The sole report on the in vivo function of a group 4 SUT concerns StSUT4 from potato (Chincinska et al., 2008). StSUT4 localized to both the plasma membrane and the endomembranes surrounding the nucleus in tobacco and potato leaves, but not to the tonoplast. This suggests that not all group 4 SUTs function in the tonoplast and that another SUT in potato may function to transport Suc between the vacuole and cytoplasm. Antisense reduction of StSUT4 expression led to altered accumulation of soluble sugars in leaves and increased phloem export of Suc. These phenotypes are the opposite of those observed in antisense StSUT1 plants (Bürkle et al., 1998), suggesting that StSUT4 has a distinct biological function. Additionally, antisense StSUT4 plants flowered early, had higher tuber production, and had reduced shade avoidance. The authors proposed that StSUT4 may act in the interplay of carbon availability and flower induction pathways.
Very little is known about other grass members of the group 4 SUTs. The rice ortholog of HvSUT2 is OsSUT2. From reverse transcription-PCR experiments, OsSUT2 is constitutively expressed in vegetative and reproductive tissues, although expression decreases toward the end of seed development (Aoki et al., 2003). Determining the physiological functions of these genes awaits the characterization of plants containing the respective loss-of-function mutations.
Similarly, the functions of group 5 SUTs are unknown and remain to be determined. OsSUT5 is expressed nearly ubiquitously and shows the highest expression level in sink leaves (Aoki et al., 2003).
FUNCTIONS OF GROUP 1 SUTs
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED Group 1 SUTs are present only in monocots and are subdivided into two clades represented by OsSUT1 and OsSUT3 (Fig. 2). Nothing is currently known of the functions of grass SUT3 genes. Here, we briefly summarize what is known about the functions of grass SUT1 genes in phloem loading in leaves as well as their roles in other tissues.
Rice
OsSUT1 was the first SUT cloned from monocots (Hirose et al., 1997). Reverse transcription-PCR shows that it is expressed at high levels in germinating seeds, source leaf sheaths, panicles, and developing grains and at lower levels in sink and source leaves (Aoki et al., 2003). OsSUT1 transcript was expressed in companion cells in leaf sheaths and in the scutellar vasculature of germinating seeds (Matsukura et al., 2000; Furbank et al., 2001; Scofield et al., 2007a). OsSUT1 may also play a role in maternal tissues, as both transcript and protein have been localized to the nucellus, vascular parenchyma tissue, and nucellar projection (Furbank et al., 2001). Promoter:GUS analyses and immunolocalization experiments showed that OsSUT1 was expressed in the companion cells and sieve elements, where it may function in phloem loading of Suc for transport to seedling shoots and roots (Scofield et al., 2007a). OsSUT1 is also expressed in the phloem along the long-distance transport path from the flag leaf blade to the base of the filling grain (Scofield et al., 2007b). It was suggested that OsSUT1 may function in Suc retrieval from the apoplasm along the transport path.
OsSUT1 does not appear to have an essential function in phloem loading of Suc in mature leaf blades. This was shown by strongly reducing OsSUT1 gene expression by antisense RNA suppression (Ishimaru et al., 2001; Scofield et al., 2002). Antisense lines with almost complete reduction in RNA or protein levels had normal vegetative growth and no alteration in photosynthesis or leaf carbohydrate contents, contrary to what would be expected if OsSUT1 principally functioned in phloem loading. Instead, transgenic plants had reduced grain filling and decreased germination, indicating that OsSUT1 plays an important role in transporting Suc to the developing grain and in remobilizing stored carbohydrates during early seedling growth. No visible phenotype was observed in leaves of OsSUT1 antisense lines, potentially due to genetic redundancy (Ishimaru et al., 2001; Aoki et al., 2003) or to utilization of a symplastic phloem-loading pathway (Kaneko et al., 1980). To our knowledge, these two reports are the only publications to date that directly assess the biological function of a grass SUT in vivo.
In support of a symplastic loading pathway in rice, fluorescent dyes were recently used to show that a xylem sap retrieval pathway functions in rice leaf blades to transfer solutes from the xylem transpiration stream into adjacent vascular parenchyma cells and into the phloem sieve elements (Botha et al., 2008). Potentially related to a functional symplastic phloem-loading pathway, grass veins contain two types of sieve elements: thick walled and thin walled (Kuo and O'Brien, 1974; Walsh, 1974; Evert et al., 1978; Botha, 2005). Botha and coworkers (2008) showed that the thick-walled sieve elements were connected by plasmodesmata to vascular parenchyma cells and accumulated significant amounts of fluorescent dye. In contrast, the thin-walled sieve elements and their companion cells were virtually symplastically isolated from other cells and contained much less dye. An intriguing possibility that needs more attention is that grass leaves may be able to use both symplastic (via thick-walled sieve elements?) and apoplastic (thin-walled sieve elements) phloem-loading mechanisms (Chonan et al., 1985; van Bel, 1993; Botha, 2005). Perhaps if the apoplastic pathway is not functional, the ability to symplastically load Suc into the phloem sustains plant growth. In fact, Srivastava et al. (2008) recently discussed the possibility that Arabidopsis may similarly be able to load Suc directly into the phloem using a completely symplastic pathway. Hence, it is possible that plants thought to use apoplastic phloem loading based on anatomical considerations are also conditional symplastic loaders (for discussion of mixed loading, see van Bel, 1993).
Wheat
In wheat, three homeologous genes known as TaSUT1A, TaSUT1B, and TaSUT1D (corresponding to the A, B, and D progenitor genomes that make up the genome of hexaploid wheat) have been characterized (Aoki et al., 2002). The three are almost equally expressed in leaves, internodes, and developing grains (Aoki et al., 2004). RNA in situ hybridization showed that the TaSUT1 RNA accumulated specifically in companion cells in leaves. However, immunolocalization experiments with an antibody raised against the rice OsSUT1 protein determined that TaSUT1 is localized to the plasma membrane of sieve elements in both leaves and stem tissues. This suggests that phloem loading of Suc occurs in the sieve elements in wheat and that the protein trafficks through plasmodesmata (Aoki et al., 2004). Based on its expression pattern, TaSUT1 has been proposed to function in phloem loading in source leaves and in retrieval of leaked Suc along the transport phloem in sheath and stem tissues. In developing grains, TaSUT1 localized to the plasma membrane of modified aleurone and subaleurone cells adjacent to the endosperm cavity, where it has been proposed to play a role in Suc uptake into filial tissues (Bagnall et al., 2000; Aoki et al., 2006).
Barley
HvSUT1 in barley is expressed at very high levels in developing seeds during the time of starch deposition (Weschke et al., 2000). RNA in situ hybridization determined that HvSUT1 transcript mainly accumulated in the maternal nucellar projection and in the endosperm transfer layer, which correspond to the site of Suc exchange between maternal and filial tissues. HvSUT1 is proposed to function in developing seeds in Suc uptake for delivery to starchy endosperm cells and in Suc retrieval in maternal tissue. HvSUT1 is expressed at lower levels in sink and source leaves, but its function in these tissues is not known. HvSUT1 RNA has been detected in phloem sap, indicating that the RNA trafficked through plasmodesmata from the companion cells to sieve elements (Doering-Saad et al., 2002). Expression of HvSUT1 in Xenopus oocytes determined that the transporter had moderate Suc affinity and was more substrate selective than AtSUC2 (Sivitz et al., 2005).
Sugarcane
Sugarcane internodes have the remarkable ability to store massive amounts of Suc, depositing it in both the vacuoles of stem storage parenchyma cells and the apoplast surrounding these cells (Welbaum and Meinzer, 1990). To prevent apoplastic backflow of Suc to the phloem, the stem veins are surrounded by sclerenchyma cells, which contain suberin and lignin in their cell walls (Walsh et al., 2005). This produces an apoplastic barrier to solute movement, as shown by the fact that xylem sap contains no Suc (Welbaum et al., 1992). Therefore, Suc must take an entirely symplastic route from the phloem into the storage parenchyma cells, where it is exported to the apoplast. Fluorescent dye tracer studies support this transport path (Rae et al., 2005; Walsh et al., 2005).
ShSUT1 RNA is expressed at the highest levels in tissues experiencing high sugar flux, specifically, mature exporting leaves and sugar-accumulating internodes (Rae et al., 2005). Immunolocalization experiments with an antibody raised against a ShSUT1 peptide indicated that the protein is not expressed in the phloem and therefore does not function in phloem loading or retrieval in these tissues (Rae et al., 2005). Instead, ShSUT1 localized to vascular parenchyma and mestome sheath cells (inner bundle sheath cell layer) in leaf and stem tissues. It was proposed that ShSUT1 may function as a biochemical barrier to apoplastic transport of Suc and retrieve any Suc leaked to the apoplast (Rae et al., 2005). Consistent with this idea, electrophysiological studies of ShSUT1 in oocytes determined that ShSUT1 had the highest selectivity for Suc of any SUT known, along with a relatively low Suc affinity (Reinders et al., 2006). It is likely that additional SUTs function in phloem loading of Suc in sugarcane, as an antibody raised against rice OsSUT1 immunolocalized to phloem tissue in leaves and stem (Rae et al., 2005). Further studies are needed to characterize additional sugarcane SUTs to determine which family member(s) function in the phloem.
Maize
ZmSUT1 is highly expressed in photosynthetic tissues, with maximal expression in leaf blades at the end of the day and minimal expression during the night (Aoki et al., 1999). Additionally, in an expanding leaf, ZmSUT1 is expressed in a gradient, with highest levels in mature tissues at the tip and lowest levels in the immature base, a pattern reflective of the sink-to-source transition. Based on its expression pattern and biochemical activity, it has been proposed that ZmSUT1 functions in phloem loading in source tissues (Aoki et al., 1999; Carpaneto et al., 2005). However, as the closely related genes OsSUT1 and ShSUT1 (Fig. 2) apparently do not function in phloem loading, it is not certain what role ZmSUT1 plays in planta. Complicating the matter, ZmSUT1 has also been shown to be capable of transporting Suc across the plasma membrane in both directions in Xenopus oocytes if the membrane potential, Suc, and pH gradients are reversed (Carpaneto et al., 2005).
We recently determined that ZmSUT1 functions in phloem loading by characterizing a knockout mutation (Slewinski et al., 2009). Zmsut1 mutant plants developed chlorotic leaves that hyperaccumulated carbohydrates and prematurely senesced. Application of [14C]Suc to abraded leaves demonstrated that Suc export was greatly diminished in Zmsut1 mutants compared to wild type. In addition, mutant plants had reduced plant height, fewer leaves, delayed flowering, and stunted tassel development. Presumably, these phenotypes result from a reduction in assimilates delivered to sink tissues due to the failure to export Suc from source leaves. These phenotypes are similar to those reported in dicot plants containing mutations in SUT genes that are responsible for phloem loading (Riesmeier et al., 1994; Bürkle et al., 1998; Gottwald et al., 2000; Hackel et al., 2006; Srivastava et al., 2008). Hence, it appears, at least in maize, that SUT1 function is essential for phloem loading of Suc. Determining the biological functions of the five additional SUTs in maize (Fig. 2) will require characterizing plants that harbor mutations in each gene.
Tie-dyed Loci
In addition to Sxd1 and ZmSUT1, several other maize genes that affect carbohydrate accumulation in leaves have been characterized. The tie-dyed1 (tdy1) mutant was identified by its variegated yellow and green leaf phenotype (Braun et al., 2006). tdy1 sectors violate the clonal cell lineages in maize leaves, suggesting that a mobile signal is responsible for sector formation. Phenotypic characterization of the mutant revealed that the tdy1 yellow sectors hyperaccumulated soluble sugars and starch, indicating that carbon partitioning was perturbed. Additionally, tdy1 mutants showed reduced plant stature, delayed flowering, and decreased yield, phenotypes that resemble dicot and maize SUT mutants (see above). However, because tdy1 mutants are variegated, it was proposed that the gene functions as an osmotic stress or sugar sensor to promote phloem loading, rather than acting directly to transport sugars (Braun et al., 2006). Through a clonal mosaic analysis, Tdy1 function was mapped to the middle layer of leaves, consisting of the interveinal mesophyll, bundle sheath, and vascular cells (Baker and Braun, 2007). To further dissect the function of Tdy1 in carbohydrate partitioning, epistasis analysis was used to determine that Tdy1 does not play a role in the photosynthetic cells in starch metabolism (Slewinski et al., 2008). In addition, it was shown that Tdy1 functions independently of Sxd1 in controlling carbohydrate accumulation in leaves (Ma et al., 2008). However, based on its nearly identical mutant phenotype and a dosage-sensitive genetic interaction, Tdy1 was found to function in the same pathway as Tdy2 (Baker and Braun, 2008). It was hypothesized that TDY1 and TDY2 may form a protein complex that promotes phloem loading.
To understand the function of Tdy1, we recently cloned the gene (Ma et al., 2009). Tdy1 encodes a novel transmembrane protein found only in grasses, although two conserved protein subdomains are present in monocots and dicots. RNA in situ hybridization studies determined that Tdy1 RNA was expressed exclusively in phloem cells. In fact, Tdy1 is expressed as soon as protophloem cells mature, indicating that Tdy1 may be a useful marker for early phloem cell differentiation. Monitoring symplastic solute movement with a fluorescent dye in wild-type and mutant leaves revealed that phloem loading was impaired in tdy1 mutants. Therefore, it was proposed that Tdy1 may function to promote phloem loading, potentially through modulating the activity of a SUT. Future investigations to examine the genetic and biochemical interactions between TDY1 and maize SUTs will test this hypothesis.
CONCLUDING THOUGHTS
TOPLEAF ANATOMYSUC ENTRY INTO A...SUTsFUNCTION OF GROUP 2...GROUP 3, 4, AND...FUNCTIONS OF GROUP 1...CONCLUDING THOUGHTSLITERATURE CITED Our understanding of the genetic regulation of carbon partitioning is just beginning. In addition to the genes described above, we have identified many other maize loci that regulate leaf carbohydrate accumulation. Characterization of the function of these genes and all SUT family members will open up new avenues of investigation into the control of carbon partitioning in plants. Moreover, it will afford exciting prospects for biotechnological approaches to enhance crop yield and biofuels production. As a case in point, it was recently demonstrated that the sugar content of sugarcane stems could be doubled with no apparent defect to plant growth, illustrating the potential to greatly modify carbon partitioning patterns (Wu and Birch, 2007). In addition, hyperaccumulation of carbohydrates in maize leaves redirected significant amounts of carbon to cellulose in the cell wall (Baker and Braun, 2008). These examples indicate that tremendous opportunities exist to manipulate carbon partitioning pathways in grasses. Additional research is needed to uncover the genetic pathways and control points regulating carbohydrate partitioning in plants to realize these goals.
ACKNOWLEDGMENTS We thank John Ward, two anonymous reviewers, and members of the Braun and McSteen laboratories for valuable comments that improved the manuscript. We gratefully acknowledge the Maize Genome Sequencing project and the Sorghum and Brachypodium Genome Projects of the Department of Energy Joint Genome Institute for making the draft genome sequences available prior to publication. We sincerely appreciate the efforts of Toby Kellogg and Robin Buell to organize this Focus Issue on the Grasses.
Received September 1, 2008; accepted October 19, 2008; published January 7, 2009.
FOOTNOTES 1 This work was supported by the National Research Initiative of the U.S. Department of Agriculture Cooperative State Research, Education, and Extension Service (grant no. 2008–35304–04597 to D.M.B.).
The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: David M. Braun (dbraun@psu.edu
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[C] Some figures in this article are displayed in color online but in black and white in the print edition.
www.plantphysiol.org/cgi/doi/10.1104/pp.108.129049
* Corresponding author; e-mail dbraun@psu.edu
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LITERATURE CITED
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