Reference Work Entry

Glycoscience

pp 1437-1472

Starch: Structure, Properties, Chemistry, and Enzymology

  • John F. RobytAffiliated withLaboratory of Carbohydrate Chemistry and Enzymology, Department of Biochemistry, Biophysics, and Molecular Biology, Iowa State University

Abstract

Starch is a very important and widely distributed natural product, occurring in the leaves of green plants, seeds, fruits, stems, roots, and tubers. It serves as the chemical storage form of the energy of the sun and is the primary source of energy for the organisms on the Earth. Starch is composed of two kinds of polysaccharides, amylose and amylopectin, exclusively composed of d‑glucose residues with α‑(1→4) linkages in a linear amylose and α‑(1→4) linkages and ∼5% α‑(1→6) branch linkages in amylopectin, both combined in a water‐insoluble granule that is partially crystalline and whose size, shape, and morphology are dependent on its biological source. The properties, isolation, fractionation, enzymatic degradation, biosynthesis, chemical modification, and specific methods of analysis of starch are presented.

Keywords

Amylose Amylopectin Granules Crystallinity Hydrogen bonding Hydrophobic bonding Isolation Fractionation Gelatinization Solubilization

Abstract

Starch is a very important and widely distributed natural product, occurring in the leaves of green plants, seeds, fruits, stems, roots, and tubers. It serves as the chemical storage form of the energy of the sun and is the primary source of energy for the organisms on the Earth. Starch is composed of two kinds of polysaccharides, amylose and amylopectin, exclusively composed of d‑glucose residues with α‑(1→4) linkages in a linear amylose and α‑(1→4) linkages and ∼5% α‑(1→6) branch linkages in amylopectin, both combined in a water‐insoluble granule that is partially crystalline and whose size, shape, and morphology are dependent on its biological source. The properties, isolation, fractionation, enzymatic degradation, biosynthesis, chemical modification, and specific methods of analysis of starch are presented.

Keywords

Amylose Amylopectin Granules Crystallinity Hydrogen bonding Hydrophobic bonding Isolation Fractionation Gelatinization Solubilization

Abbreviations

ADPGlc

adenosine diphosphate glucose

α‑Glc-1-P

α‑d‑glucopyranosyl-1‑phosphate

avg. d.p.

average degree of polymerization

CGTase

cyclomaltodextrin glucanyltransferase

d.s.

degree of substitution

DMSO

dimethyl sulfoxide

EDTA

ethylene diamine tetraacetic acid

FACE

fluorescence‐assisted capillary electrophoresis

Pi

inorganic phosphate

PPO

2,5-diphenyloxazole

POPOP

1,4-bis-2-(5-phenyloxazoly)benzene

TEMPO

2,2,6,6‑tetramethyl-1‑piperidine oxoammonium ion

UDP-Glc

uridine diphosphate–glucose

35.1 Introduction, History, Development, and Uses

Starch is an abundant, naturally occurring polysaccharide, rivaling cellulose in the amount found on the Earth. It is found in the leaves of all green plants and in the seeds , fruits , stems, roots , and tubers of most plants. Starch results as an end-product of photosynthesis and serves as the chemical storage form of the energy of the sun on the Earth. A high percentage of the energy available to non‐photosynthetic organisms comes from starch, which is found in the principal food crops of the world: wheat, potatoes, rice, maize, barley, rye, beans, peas, sorghum, tapioca (or cassava), sweet potatoes, avocados, arrowroot, taro, bananas, mango, pineapple, sago (palm starch) and so forth.

It is estimated that 60–70% of the caloric intake by humans comes from starch. As such, starch has been of great importance in the evolution of organisms and especially for humans, where it also has had a role in the evolution of culture. Besides being used as an essential food, wheat starch was used to give body and the ability to hold ink to papyrus, a thin bark that was the earliest material used for writing (∼4000 b.c.), especially in Egypt. When paper was developed in China, around 100 a.d., starch was also used to give body to the paper and to hold ink on the paper and it continues today to be used to size paper. The Romans (∼100 b.c.) used starch to whiten cloth and to powder hair, and about 300 a.d., it was widely used to stiffen cloth and was often mixed with dyes to color the cloth. Colored starches (especially yellow and red) were also used as cosmetics .

Starch is one of three major biorenewable materials found on the Earth, sucrose and cellulose being the other two. Of the three, starch is today the most important because of its relative abundance and its relative ease of isolation in a highly pure form, which is relatively easily solubilized and enzymatically hydrolyzed to glucose and/or different maltodextrin products, or chemically modified. It, therefore, is a biorenewable, natural product that finds many industrial uses, such as the formation of glucose syrups, high-fructose syrups, maltodextrins with a wide range of average molecular weights, cyclomaltodextrins, d‑glucitol (d‑sorbitol) and the formation of ethanol, acetic acid, d‑lactic acid, and other organic compounds by fermentation.

35.2 Isolation of Starch Granules

Because of the water‐insolubility of starch granules, they are relatively easy to isolate from their plant sources. The source, for example seeds from maize, wheat, barley, rice, beans, and so forth are first steeped in water for 10–15 h at 50 °C. Steeping softens the outer parts of the seeds so the starch inside can be more easily obtained. The steep water often contains 1.63 g of sodium hydrogen sulfite per liter to remove protein. In some cases, the sulfite is omitted to retain the protein and give a starch as it more closely exists in the plant. The steeped material is then ground, milled, or blended to give a white insoluble suspension of starch that is filtered one time through six layers of cheese cloth and a second time through eight layers, to remove fibrous non-starch substances. The starch is allowed to settle and the water is poured off and fresh water is added and the starch is washed two or three times in this manner. The starch is then centrifuged or filtered and allowed to air dry at 20 °C. It is then lightly pounded into a fine powder.

If the starch is to be isolated from a tuber, stem or rhizome, the plant materials are washed and peeled and then cut into 1-cm pieces, placed in water, and blended. The resulting starch suspension is then obtained in a similar manner, as described above, beginning with filtering through the cheese cloth.

35.3 Chemical Composition of Starch

Most starches are composed of two kinds of polysaccharides, a linear α‑(1→4) linked glucan, called amylose , and an α‑(1→4) linked glucan with 4.2 to 5.9% α‑(1→6) branch linkages, called amylopectin . See Fig. 1 for the chemical structure of a segment of amylose and a segment of amylopectin and Table 1 for the percent branching in amylopectin for several different starches.

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Figure1

The structures of segments of amylose and amylopectin

Table 1

Percent branching of amylopectin from different starches

Amylopectins

% Branchinga

Maize

4.2

Potato

4.5

Wheat

4.8

Barley

5.0

Oats

5.2

Waxy maize

5.9

aThe avg. % branching was calculated from the avg. chain length determined by the enzymatic method [177]

The ratio of amylose to amylopectin also varies, depending on the source of the starch; it ranges from 17 to 70% amylose and a corresponding 83 to 30% amylopectin. See Table 2 for a list of different starches and their composition of amylose and amylopectin. Usually amylopectin is present as the major component, with a common ratio of 1:3 or ∼25% amylose and ∼75% amylopectin. There are mutants, however, such as, waxy barley, waxy maize , waxy rice, waxy potato, and waxy sorghum that are 100% amylopectin and there are the high-amylose starches, such as, amylomaize-5 , with 53% amylose and 47% amylopectin, amylomaize-7 , with 70% amylose and 30% amylopectin, and wrinkled pea with 66% amylose and 34% amylopectin.

The molecular weights of the two polysaccharides also vary, depending on the source of the starch. Amylose is much lower than amylopectin and both polysaccharides exist as a distribution of molecular weights, so that the determined molecular weight is an average. Amylomaize-7 amylose has an average degree of polymerization (avg. d.p.) of ∼400 glucose residues or a number average molecular weight of ∼64,800 Da; potato amylose has an avg. d.p. of ∼1,000 glucose residues or a number average molecular weight of ∼162,000 Da; and wheat amylose has an avg. d.p., of ∼4,000 glucose residues or a number average molecular weight of 648,000 Da. Amylopectin is a much larger molecule, having number average molecular weights of 1 × 106 to 5 × 108 Da. The polydispersity of the sizes of the amylose component is lower than the amylopectin component.

Table 2

Composition of starch granules from ten plant sources

Starch

Amylosea(%)

Amylopectina(%)

Lipidb(%)

Proteinb(%)

Phosphateb(%)

Maize

25

 75

0.80

0.35

0.090

Waxy maize

 0

100

0.20

0.25

0.024

Amylomaize-5

53

 47

0.70

0.30

0.090

Amylomaize-7

70

 30

0.75

0.30

0.060

Potato

22

 78

0.01

0.10

0.210

Wheat

23

 77

0.90

0.40

0.180

Rice

19

 81

0.59

0.30

0.090

Tapioca

17

 83

0.02

0.10

0.009

Banana

20

 80

0.48

0.32

0.060

Shoti

30

 70

0.01

0.20

0.630

aThe percent amylose and amylopectin were determined by precipitating the amylose with 1‑butanol and then precipitating the amylopectin with two volumes of ethanol; the butanol-1 was removed and both were dehydrated by trituration with acetone several times and ethanol, followed by drying under vacuum at 40 °C, and weighed.bFrom Ref. [178]

It has been determined that for a starch granule of 15 µm diameter and a mass of 2.65 × 10−9 g, containing 25% amylose, with an avg. d.p. of 1,000 and 75% amylopectin, with an avg. d.p. of 100,000, there would be 2.5 × 109 molecules of amylose and 7.4 × 107 molecules of amylopectin in a granule [1].

Two minor carbohydrate components have been found in starch granules. The first is a water-soluble amylose that is slightly branched ∼0.5 to ∼2.0% [2,3,4]. The second is a very highly branched component found in 12 varieties of starch granules that were isolated and examined for their structure and properties and most probably occur in all starches. They ranged in amounts from 0.51% (w/w) in amylomaize-7 starch to 8.4% (w/w) in rice starch. They had relatively high molecular weights, 2.47 kDa for amylomaize-7 starch to 5.75 kDa for waxy maize starch, and a high degree of α‑(1→6) branching, ranging from 15.6% for rice starch to 41.4% for shoti starch [5].

In addition to these major carbohydrate components, starch granules also contain minor non‐carbohydrate components: lipids from 0.01 to 0.80% (w/w); proteins, from 0.10 to 0.40% (w/w); and phosphate 0.09 to 0.63% (w/w). See Table 2 for the compositions of ten types of starch granules. The lipids are primarily found in the cereal starches: maize, waxy maize, amylomaize-5, amylomaize-7, rice, wheat, barley, rye, and so forth; and the phosphate in these starches are part of the phospholipids. The phosphate in the tuber starches, canna, potato, shoti and others, is attached to the primary alcohol of the glucose residues in amylopectin. At least some of the protein, if not all of it, is the enzymes, starch synthases and starch branching enzymes that were responsible for the biosynthesis of the starch in the granule (see Sect. 8). In addition to the composition mentioned above, starch granules at equilibrium with their environment (20 °C) and humidity (40–50%) will absorb water and contain 10–15% (w/w) water of hydration. When starch granules are suspended in water, they swell to a limited extent and adsorb water to ∼30% (w/w).

35.4 Occurrence of Starch as Water-Insoluble Granules

All starches are stored in plants as water‐insoluble particles or granules in the chloroplasts in leaves and in amyloplasts in other plant tissues. The granules are relatively dense particles of compact molecules, with semi‐crystalline properties. They were first observed in the early eighteenth century by van Leeuwenhoek, using his newly invented microscope. In the middle of the nineteenth century, Nägeli [6] described differences in the size and shapes of starch granules from various plant sources. In 1913, Reichert [7] published a book containing hundreds of photomicrographs of starch granules from widely different plant sources. In 1994, Jane et al. [8] published a comparative anthology of scanning electron micrographs of starch granules from 54 plant sources, showing widely different sizes of large (100 µm) to very small (0.2 µm) granules and different morphologies, such as smooth, oval, and spherical shapes, round pancake shapes with varying thicknesses, flat thin plates or lenticular‐shaped disks, and irregularly shaped polygons whose edges have varying degrees of sharpness. Canna bulb starch granules are very large (60 × 100 µm) and potato starch granules are also large (20 × 75 µm). Many of the starches from the cereal grains, such as maize, waxy maize, oats, and sorghum are smaller (15–25 µm), with the exception of rice starch granules, which are quite small (3–5 µm). The cereal starches have irregular polygonal shapes, with a number of faces that have relatively sharp edges. Starches from beans and peas have smooth oval granules, with 10–45 µm diameters. They often are accompanied by an indentation in the center or at the end of the granule. There are some starches that are extremely small, such as the granules from Chinese taro (1–4 µm), amaranth (0.5–2 µm), and parsnip (1–3 µm); they generally have polygonal shapes, with sharp edges, similar to the cereal starches, but much smaller. Most leaf starches are very small, biconvex granules of ∼1 µm in diameter. Figure2 illustrates these differences by pictures of scanning electron micrographs of nine starch granules with varying sizes and morphologies.

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Figure2

Scanning electron micrographs of starch granules from nine different botanical sources: A, potato; B, rice; C, wheat; D, mung bean; E, maize; F, waxy maize; G, tapioca; H, shoti; J, leaf starch. Magnification 1500X for each starch

35.4.1 Structure of the Starch Granules

Starch granules also have varying degrees of crystallinity, as shown by X‑ray powder pattern diffractions. Three distinct X‑ray patterns have been observed, A-, B-, and C‑patterns [9,10]. The A‑patterns are characteristic of cereal grain starches, such as maize, waxy maize, wheat, and rice. The B‑patterns are characteristic of tuber, fruit, and stem starches, such as canna, potato, sago, banana starches and the mutant maize starches, amylomaize‑5 and amylomaize‑7 [9,10]. C‑type patterns found in roots, beans, and peas are intermediate between A- and B‑types [10,11]. Tapioca starch gives a typical C‑type X‑ray pattern. It has now been shown that C‑type starch granules have a central core with a B‑type structure, surrounded by an A‑type structure [12].

When B- and C‑type native starches are heat-moisture treated, 30 °C for B‑type and ∼50 °C for C‑type, they are converted into A‑type X‑ray patterns [13,14,15,16]. Conversion of A‑type starches into other crystalline forms only occurs when the original structure is completely destroyed and it is then allowed to recrystallize [17].

The interpretation of the X‑ray diffraction patterns of starch granules and the significance of the A- and B‑types of granules are problematic. There are differences in the ultrastructures of starch granules from different plant sources [18]. This was reinforced by a study of the kinetics and products of the reaction of glucoamylase with seven native starch granules in which three distinct categories were obtained: waxy maize starch was the most susceptible; barley, maize, and tapioca starches were intermediate; and amylomaize‑7, shoti, and potato starches were quite resistant [19].

A general model for the crystalline parts of the starch granule has developed as a spherocrystalline assembly of disk-shaped amylopectin molecules [20] that are radially directed with the non-reducing-ends of the chains directed toward the surface of the granule [21,22]. The amylose chains in the cereal starches exist as lipid complexes that are left-handed extended helices, arranged tangentially in the granule. For non‐complexed amylose chains in the cereal starches and in other starches, the amylose chains exist as both single and double helices [23,24]. In the granule, the chains of the amylopectin molecules can be considered as two‐dimensional molecules that stack on top of each other, along with amylose chains, to make a three‐dimensional micelle, giving radially oriented crystallites 9 nm in size. These units of both crystalline and amorphous phases form a network, extending throughout the granule in repeating units that are a common structural feature for all starches [25]. The amylose and amylopectin molecules form hydrogen bonds and hydrophobic bonds , both intermolecularly and intramolecularly, to varying degrees that hold the molecules together, giving a water‐insoluble granule. The arrangement and quantity of these bonds in the granules gives rise to the differences in granule properties (degrees of crystallinity, types of X‑ray diffraction patterns , gelatinization temperature, and susceptibility to enzyme hydrolysis [19,26,27]) found in starches from different plant sources. The α‑(1→6) branch linkages in amylopectin are believed to occur in the amorphous regions of the granule. The branch linkages of amylopectin are clustered, with several chains of 15–20 glucose residues from the cluster, intertwined into double helices, contributing to the crystallinity [17,21].

35.4.2 Growth Rings in Starch Granules

Growth rings are readily seen in the large tuber starch granules, such as potato or canna starches, by optical microscopy. Growth rings are particularly apparent by scanning electron microscopy of granules that have been treated with acid or α‑amylases [9]. Figure3 is a scanning electron micrograph of a potato starch granule that has been treated with Bacillus amyloliquefaciens α‑amylase. During treatment with α‑amylase, the amorphous layers in the granule are hydrolyzed and the crystalline layers remain intact. These so-called growth rings most likely originate during different phases of the biosynthesis of starch granules in which the deposition of crystalline layers of starch alternates with an amorphous layer of starch. These layers most probably are produced by fluctuations in the rate and/or mode of starch deposition during biosynthesis. Potato and canna starch granules show the growth rings the best when they are fully hydrated. When the starch granules are dried, the growth rings disappear [28].

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Figure3

Scanning electron micrograph of potato starch granule treated with Bacillus amyloliquefaciens α‑amylase, showing growth rings of the crystalline α‑amylase resistant starch in the granule

35.4.3 Gelatinization of Starch Granules and the Solubilization of the Starch

The first step in the utilization of starch often involves the disruption of the granule, although by no means is this always the case (see Sect. 9 on the chemical modification of starch granules). Disruption of starch granules can be accomplished by heating a suspension of starch granules in water, either by boiling the water (100 °C) or by autoclaving the suspension under pressure at 121 °C.

Starch granules swell when heated in water and have a “gelatinization temperature ” that is characteristic of their source. When this temperature is reached, the swollen granules loose their birefringence , as evidenced by the loss of the “polarization cross” that can be observed with a polarizing microscope . The granules, however, still have a certain amount of structural integrity, but as the temperature is increased further, they continue to swell and eventually burst, releasing the amylose and amylopectin molecules, along with the minor components, into the aqueous solution. The heating process promotes increased motion of the molecules in the granule, eventually breaking the hydrogen and hydrophobic bonds between the molecules in the crystalline regions of the granule, which become hydrated and are released into the water.

Cold, chemical methods of solubilization of starch can also be used, such as alkali (1 M NaOH) and dimethyl sulfoxide (DMSO) that gelatinize starch granules at low temperatures (20 °C). Gelatinization and the consequent solubilization of starch granules in DMSO occurs by a very different path from gelatinization in boiling water or autoclaving. Starch granules do not swell in DMSO as they do in boiling water [29,30], instead the granules slowly dissolve in anhydrous DMSO by fragmentation of the interior of the granules into smaller and smaller pieces. The addition of water to DMSO, however, greatly increases the rate of gelatinization and alters the pathway, which is different from gelatinization in anhydrous DMSO. Potato and canna starches retain their shape and birefringence during dissolution in aqueous DMSO and fragmentation does not occur. Instead the starch is peeled from the surface of the granules, with large pieces of starch gel adhering to the smooth surfaces of the ungelatinized portions of the granule [31]. When water is added, the DMSO breaks hydrogen bonds of the starch molecules at the surface of the granule and the water hydrates and gelatinizes the starch molecules at the surface, peeling the starch from the granules in layers [32].

Another method of chemically gelatinizing starch granules is by the use of relatively high concentrations of salt solutions to produce swelling and dissolution [33]; 4‑M CaCl2 has been used to gelatinize potato starch granules [34] and 13‑M LiCl to gelatinize the starch from the surface of maize starch granules [35]. A method has been developed in which the surfaces of eight starch granules were peeled under controlled conditions, using aqueous-DMSO to give slow, gradual peeling from the surface of the granules in 8–10 h by selecting the amount of water from 5 to 15% (v/v) and the temperature from 0–15 °C [36].

There is evidence from two sources that the gelatinization process does not completely disrupt the hydrogen and hydrophobic bonds between the starch molecules, even though they are dissolved in the water. The first is the significant and greatly different activities of a single kind of α‑amylase hydrolyzing solubilized starch from different plant sources [26]. The second is the differences that are observed in the water‐solubility of starches from different plant sources and from acid/alcohol modified starches [32]. The maximum water solubilities for seven common starches are given in Table 3.

Table 3

Water solubility of seven native starches by autoclaving 220 mg in 10 mL at 121 °C for 20 min

Starches

Solubility (mg/mL)

Waxy maize

18.7

Tapioca

17.4

Potato

12.4

Maize

12.4

Rice

 7.9

Amylomaize-7

 5.5

Wheat

 5.2

From Ref. [32]

35.5 Fractionation of Starch into Amylose and Amylopectin

Early methods in 1940 for fractionating starch involved the leaching of an amylose component from swollen starch granules with hot water [37,38,39,40]. Starch granules (1–2% w/v) were successively treated with 70 °C water and the linear amylose component was obtained. This extraction, however, was found to be incomplete [41] and only the low molecular weight amylose was extracted. A few years later, the amylose and amylopectin components were completely separated by first solubilizing the starch granules in boiling water and selectively precipitating the amylose from the solution by 1‑butanol  [41,42]. This 1‑butanol complex was crystalline, giving a V‑type X‑ray pattern  [43]. After the removal of the 1‑butanol‐amylose precipitate by centrifugation, the amylopectin component was precipitated from the supernatant with two volumes of anhydrous ethanol. The 1‑butanol can be removed from the amylose and the two components can be dehydrated by being treated several times with anhydrous acetone and once with anhydrous ethanol, followed by drying under vacuum at 40 °C to give dry powders. It is emphasized that the lipids complexed in the granules of the cereal starches must first be removed by defatting with methanol, ethanol, or 2‑methoxyethanol (methyl cellosolve) extraction before solubilizing the granule and removing the amylose by adding an organic complexing agent [44], otherwise the removal of the amylose in these starches is incomplete.

Table 4

Hydrophobic compounds forming complexes with amylose and those that do not form complexes

Compounds forming complexes

1,1,2,2‑tetrachloroethane

bromocyclopentae

cycloheptane

1,2‑dichloropropane

hexachloroethane

chloroform

(1.2.2)-bicycloheptene

fluorobenzene

1,1,2,2‑tetrabromomethane

1,1,2‑trichloroethane

cyclohexane

cyclopentane

carbontetrachloride

methylcyclohexane

cyclohexene

methylcyclopentane

cycloheptene

2,3‑dibromobutane

cycloheptadiene

2,3‑dimethylbutane

1,1,1‑trichloroethane

thymol

Compounds that do not form complexes

tetrachloroethylene

carbon disulfide

trichloroethylene

n-heptane

chlorobenzene

mineral oil

bromobenzene

1‑bromopentane

iodobenzene

1‑bromobutane

p-cymene

p-dibromobenzene

toluene

p-chlorobromobenzene

m-xylene

1,3,5‑triethylbenzene

p-xylene

1‑methylnaphthalene

mesitylene

 

From Ref. [46]

Another organic compound that forms a complex with amylose is thymol, which also has been used to separate amylose from amylopectin [45]. Thymol is added to a starch solution as a 10% (w/v) solution in ethanol. Besides 1‑butanol and thymol, many other organic compounds have been found to form complexes with amylose and give precipitates [46]. A list of organic compounds that form complexes with amylose and a list of those that do not form a complex and precipitate amylose are given in Table 4.

35.6 Properties of Amylose and Amylopectin

It has been known for almost 200 years that starch gives a deep blue color when a solution of potassium iodide and iodine is added [47]. More than a century later it was suggested that the complex consisted of a helical polysaccharide, with triiodide in the center of the helix [48]. Using flow dichroism, it was demonstrated that the triiodide was stacked in a linear structure, as required for the helical model [49]. Another study of the optical properties of crystals of the amylose‐triiodide complex showed it to be consistent with a helical structure [50] and X‑ray diffraction showed the triiodide complex gave the dimensions of a unit-cell of a helix with six glucose residues per turn [51]. This confirmed a helical structure for the amylose complex with triiodide that predated the helical models proposed by Pauling for polypeptides [52] and the double helical model for DNA by Watson and Crick [53] by 10 years.

The amylose‐organic compound complexes give a particular kind of X‑ray pattern that is called a “V‑pattern” [54,55,56]. Amylose complexed with 1‑butanol or 1‑hexanol forms helices with six glucose residues per turn of the helix; branched chains or halogenated alkanes, such as tert-butyl alcohol (2‑methyl-2‑propanol) and tetrachloroethane form helices with seven glucose residues per turn of the helix [46,57,58,59], and bulkier molecules, such as 1‑naphthol and thymol form helices with eight glucose residues per turn of the helix [60]. The length of the helical chain that is folded back and forth, however, remains at 10 nm for all of the complexing agents.

α‑Amylases hydrolyze the amorphous, folding areas on the surfaces of the lamella of packed helices, with the formation of resistant amylodextrin helical complexes of different degrees of polymerization (d.p.) values that are dependent on the number of glucose residues per turn of the helix: 1‑butanol complex (six glucose residues per turn) gave d.p. 75 ± 4, tert-butyl alcohol and 1,1,2,2‑tetrachloroethane (seven glucose residues per turn) gave d.p. 90 ± 3, and 1‑naphthol (eight glucose residues per turn) gave d.p. 123 ± 2 [61]. See Fig. 4 for the formation of these amylodextrin complexes by α‑amylase hydrolysis of the surface of the folded amylose‐organic complex.

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Figure4

Alpha-amylase hydrolysis of the organic molecule (1‑butanol, tetrachloroethane, or 1‑naphthol) amylose complexes of the lamella‐crystalline material in which the α‑amylase hydrolysis takes place at the tops and bottoms (arrows) of the complexes, giving a resistant amylose‐complex fragment of approximately the same size with avg. d.p. values of 108, 122, and 169, respectively, for the three organic complexes. From ref. [61], by permission of the authors and the publisher, Elsevier Press

The reason that amylose forms a complex with the organic molecules is that it is a relatively long, linear molecule, whose structural features inside the helix are hydrophobic, allowing hydrophobic bonding with the organic molecules, whereas amylopectin chains are much shorter because of branching and do not form a sufficient length of helical complex with organic compounds that can give the lamella crystalline complexes, thus, allowing amylose to be separated from amylopectin.

It is the amylose component of starch that gives the blue color when KI/I2 solution is added. To study the iodine‐iodide color of amyloses of different d.p. values, maltodextrin‐amylose molecules, with various avg. d.p. values from 6 to 568 were prepared by Bailey and Whelan [62], using phosphorylase, α‑d‑glucopyranosyl‐1‑phosphate, and maltohexaose. The colors of the various sized maltodextrins (1 mg) were observed when 10:1 (w/w) KI/I2 solution was added. The first color to be observed was faint red for avg. d.p. 12; a red-purple color was observed for avg. d.p. 31; a purple color was observed for avg. d.p. 40; and a blue color was observed for avg. d.p. 45. The increase in the “blue value ” was linear as a function of avg. d.p. up to ∼ avg. d.p. 60; the absorbance at 645 nm then slowly increased and reached a maximum at avg. d.p. of 400. The intensity of the iodine/iodide color in the low molecular weight range was dependent on the concentration of the iodine. When the concentration of the iodine was increased 10‑fold, the intensity was increased 50% [62].

Amylose in a water solution is thought to behave as a random coil that has some helical character [63]. The random coiled amylose moves randomly in the aqueous solution with helices constantly being formed and deformed, giving a dynamic structure. An aqueous solution of amylose, 2 mg mL−1 or more is unstable and comes out of solution (retrogrades), as an insoluble precipitate, at a rate that is dependent on the concentration. At a concentration of 1 mg mL−1, however, this same amylose does not retrograde and stays in solution indefinitely.

Acid hydrolysis of retrograded amylose (avg. d.p. 1,000) gives a resistant fragment with an avg. d.p. of 32 [61]. Hydrolysis of the retrograded amylose by human salivary α‑amylase and porcine pancreatic α‑amylase also gives a resistant fragment of avg. d.p. of 43, and hydrolysis by Bacillus amyloliquefaciens α‑amylase gives a resistant fragment of avg. d.p. of 50 [61]. A structure for retrograded amylose was proposed in which crystalline double helical chains of 32 glucose residues of ∼10 nm lengths are interspersed with amorphous regions to give resistant double helical structures. When the retrograded amylose is acid hydrolyzed for prolonged periods, most of the α‑(1→4) glycosidic linkages in the amorphous regions are hydrolyzed, leaving the double helical, crystalline regions of 32 glucose residues. When the retrograded amylose, however, is hydrolyzed by α‑amylases, they cannot hydrolyze the α‑(1→4) glycosidic linkages close to the crystalline regions because of the numbers of their substrate‐subsites, leaving stubs of glucose residues on the ends of the retrograded amylose chains, giving larger fragments than acid hydrolysis. Porcine pancreatic α‑amylase has five subsites and human salivary α‑amylase has six subsites, giving a chain with ∼10–11 more glucose residues, 32 + 11, or 43 glucose residues, and B. amyloliquefaciens α‑amylase has nine subsites, giving a chain with 18 more glucose residues than acid hydrolysis, nine glucose residues on each end, giving a total of 32 + 18 or 50 glucose residues [61].

Amylopectin acts very differently in aqueous solution. Even though it is 3- to 5‑orders of magnitude larger than amylose, it is much more water soluble. This is primarily due to the presence of ∼5% α‑(1→6) branch linkages in the amylopectin molecule. The branch linkages of 50 for every 1,000 linkages give an average chain length of 20 glucose residues per chain. Because of the branching, the amylopectin chains are, thus, relatively short and, therefore, cannot line up in solution to give intermolecular double helices of sufficient length and they, hence, do not give a significant mixture of crystalline and amorphous regions in solution that retrograde. This then is the reason why amylopectin is much more soluble and stable in an aqueous solution than is amylose.

The amylopectin molecules are not compact or spherical as are glycogen molecules, which have 10–12% α‑(1→6) branch linkages, randomly distributed. Amylopectin has relatively high viscosity, indicating an elongated structure. Electron microscopy, X‑ray diffraction, chemical, and enzymological studies have shown that amylopectin has its branch linkages clustered at ∼70 Å intervals [64]. Amylopectin molecules are ∼100–150 Å in width and ∼2,000–4,000 Å in length. Further, there is a regular alternation of elongated α‑(1→4) linked chains and clustered α‑(1→6) branch linkages that gives the 70 Å periodicity of the clustered branch linkages [64]. The amylopectin molecules can be considered to be two‐dimensional, with the third dimension coming from stacking of the molecules through the formation of intermolecular hydrogen and hydrophobic bonds in the water‐insoluble granule and also partially when in an aqueous solution [26,32]. See Fig. 5 for a schematic drawing of amylopectin and its clustered branches and its branched chains.

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Figure5

Schematic picture (looking from the top down) on a segment of amylopectin, showing the clustering of the α‑(1→6) branch linkages in a repeating sequence

As was mentioned previously, the crystalline regions in the starch granules are primarily made up of amylopectin molecules, but with the amorphous regions being made up of the clustered branch linkages of amylopectin. The amylose molecule has one reducing-end and one non-reducing-end, and the amylopectin molecule also has one reducing-end, but many non-reducing-ends due to the branching.

Retrograded and dehydrated amylose is very water insoluble. It can, however, be put into an aqueous solution by first dissolving ∼100 mg in 1 mL of DMSO by stirring and if necessary gently warming to 40–50 °C. After the amylose is completely dissolved in the DMSO, a thick clear gel-like solution is obtained. This gel can then be very slowly diluted with water, while stirring, to give the desired concentration of amylose in water. Amylopectin is readily dissolved in boiling water to give solutions of 2 to 5% (w/v).

35.7 Enzymatic Degradation of Starch

Several different kinds of enzymes degrade starch to give different kinds of products.

35.7.1 Endo-Acting Alpha-Amylase Hydrolysis of Solubilized Starch

The major category of enzymes that hydrolyze the α‑(1→4) linkages of starch is the α‑amylases . These enzymes are ubiquitous and are produced by bacteria, fungi, plants, and animals. In mammals, there are two specific sources, the salivary glands that secrete the enzyme into the mouth and the pancreas that secretes the enzyme into the small intestine. The α‑amylases are endo-acting enzymes that attack the interior parts of the polymeric starch chains, producing a rapid drop in viscosity and a rapid change in the iodine‐iodide color; because of these properties, they are sometimes called liquefying enzymes . When α‑amylases encounter a starch chain and hydrolyze an α‑(1→4) linkage, they also produce low molecular weight maltodextrins by a process called multiple attack on one of the two chains that were initially cleaved [65,66]. A common misconception about the action of α‑amylases is that they act randomly, but this is not true. The major maltodextrin products that are formed are of specific sizes and depend on the particular biological source of the α‑amylase and they, therefore, do not act randomly.

Human salivary and porcine pancreatic α‑amylases primarily produce maltose , maltotriose , maltotetraose  [67,68], and several kinds of α‑(1→6) branched maltodextrins, ranging in size from four to seven glucose residues [69,70]. An even more common misconception about the action of α‑amylases is that glucose is the major product. Glucose, in fact, is only a very minor product, being produced by a slow secondary hydrolysis of the primary maltodextrin products. For example, salivary and pancreatic α‑amylases slowly hydrolyzes maltotetraose to two molecules of maltose, the primary reaction; they also more slowly hydrolyze maltotetraose to give glucose and maltotriose and they hydrolyze maltotriose even more slowly to glucose and maltose; and they do not hydrolyze maltose at all [67,68].

A common bacterial α‑amylase is excreted by Bacillus subtilis (whose name later was changed to B. amyloliquefaciens). It primarily produces maltotriose, maltohexaose, and maltoheptaose as the major products [71]. It also only produces glucose from a slow secondary hydrolysis of the primary maltodextrin products; it very slowly hydrolyzes maltohexaose to glucose and maltopentaose; it slowly hydrolyzes maltoheptaose to maltose and maltopentaose, and it slowly hydrolyzes maltoheptaose to maltohexaose and glucose; maltopentaose and smaller maltodextrins are not hydrolyzed [71]. Another α‑amylase that is very similar to B. amyloliquefaciens α‑amylase is the plant enzyme, barley malt α‑amylase  [72]. The bacterial α‑amylase from B. licheniformis gives high yields (33%) of maltopentaose as a primary product [73]. It has a high temperature optimum of 70 ℃ and a broad pH optimum from 5 to 8, with significant activity in the alkaline pH range of 9–10 [74].

35.7.2 Exo-Acting Beta-Amylase and Glucoamylase Hydrolysis of Solubilized Starch

Starch exo-acting enzymes act exclusively at the non-reducing-ends of starch chains to form maltodextrin or glucose products. A primary exo-acting amylase is beta-amylase that is almost exclusively found in plants. The major β‑amylases that have been studied are from sweet potato, barley, and soy beans [75]. β‑Amylases catalyze the hydrolysis of the penultimate glycosidic bond at the non-reducing-ends of starch chains, forming β‑maltose. Amylose is completely hydrolyzed; hydrolysis of amylopectin, however, only gives 55% β‑maltose because the action of the enzyme is blocked by the α‑(1→6) branch linkages that β‑amylase cannot hydrolyze. This limitation leaves ∼45% of a high molecular weight limit dextrin that contains all of the branch linkages of the original amylopectin molecules at the non-reducing-ends of the limit dextrins [76]. The first β‑amylase from a source other than plants was reported from B. polymyxa [77]. It acted identically to the plant β‑amylases. An α‑amylase was reported from Streptomyces sp. that is an endo-acting enzyme, but produced 75–80% α‑maltose [78].

Glucoamylase is another exo-acting enzyme that hydrolyzes the first glycosidic linkage at the non-reducing-ends of starch chains to give β‑d‑glucose. Many fungi, such as Aspergillus niger , A. awamori, Rhizopus delemar , and R. niveus elaborate glucoamylases [79,80,81,82,83]. In contrast to other types of amylases, glucoamylase can hydrolyze both α‑(1→4) and α‑(1→6) glycosidic linkages, although at different rates, with the α‑(1→4) linkages being hydrolyzed ∼600‑times faster than the α‑(1→6) linkages. But because of the hydrolysis of the α‑(1→6) branch linkages, glucoamylases can completely hydrolyze all of the starch to β‑d‑glucose.

Another exo-acting starch‐degrading enzyme is phosphorylase . It is produced by many plants and may be a primary enzyme involved in starch degradation in plants, although many plants also have α‑ and β‑amylases. Phosphorylase, however, is not a hydrolase and catalyzes the reaction between the α‑(1→4) linkage of the terminal glucose residue at the non-reducing-ends of starch chains and inorganic phosphate to give α‑d‑glucose‑1‑phosphate [84]. Phosphorylase, like β‑amylase, is unable to by-pass α‑(1→6) branch linkages. It does not even remove all of the glucose residues close to the branch linkages and stops four glucose residues from the branch point on the two chains. Another enzyme, starch debranching enzyme, is required to hydrolyze the α‑(1→6) branch linkages for further cleavage of the α‑(1→4) linkages by phosphorylase.

Pseudomonas stutzeri produces an enzyme that forms maltotetraose from the non-reducing-ends of starch chains and, like β‑amylase, cannot bypass the α‑(1→6) branch linkages of amylopectin [85]. It gives 42% (w/w) α‑maltotetraose and 58% limit dextrin. A similar exo-acting amylase is elaborated by Aerobacter aerogenes (syn: Klebsiella pneumoniae subsp. aerogenes; Enterobacter aerogenes) and exclusively forms maltohexaose from starch [86,87].

Another exo-acting starch enzyme forms cyclomaltodextrins . The enzyme is not a hydrolase, but a glucanyltransferase and is called cyclomaltodextrin glucanyltransferase (CGTase) . The C4–OH group of the non-reducing glucose residue of starch chains is made to attack C1 of the sixth, seventh, and eighth α‑(1→4) glycosidic bonds from the non-reducing-end to form cyclic, non-reducing α‑(1→4) linked maltodextrins, containing six, seven, and eight glucose residues, respectively. Several different enzymes of this type are elaborated by different species of bacteria. The first CGTase to be isolated was from Bacillus macerans and it primarily forms cyclomaltohexaose, with smaller amounts of cyclomaltoheptaose and cyclomaltooctaose [88]. CGTases are also elaborated by B. megaterium [89] and B. circulans [90] that primarily form cyclomaltoheptaose , and a Brevibacterium sp. CGTase that primarily forms cyclomaltooctaose  [91].

Isoamylases that exclusively hydrolyze the α‑(1→6) branch linkages of starch were first recognized in plants and first isolated from broad beans [92]. A bacterial isoamylase was obtained from the culturing of Pseudomonas amyloderamosa  [93,94,95,96] and has found wide use in studying the structure of amylopectin and related polysaccharides [97].

A new exo-acting enzyme, a starch lyase, has been discovered in red seaweeds and fungi. It removes glucose residues at the non-reducing-ends of starch chains and converts them into 1,5‑anhydro‑d–fructopyranose units. This product has been found to be an antioxidant and a new and versatile chiral building block for the formation of fine chemicals [98,99].

35.7.3 Reaction of Enzymes with Starch Granules

Starch granules generally have been thought to be resistant to hydrolysis by amylases [100], although as early as 1879 [101], they were reported to be digested by amylases. Ten percent hydrolysis by α‑amylases of starch granules were reported [102]. In a study of the hydrolysis of starch granules by α‑amylases from four sources (malt, fungi, bacteria, and pancreas), it was found that there were differences, with pancreatic α‑amylase being the most effective, followed by malt, bacterial, and fungal α‑amylases [103]. The activity of B. subtilis var. amyloliquefaciens varied according to the type of starch [104], with waxy maize being the most susceptible and high amylose starch (amylomaize‑7 starch) the least susceptible.

The hydrolysis of starch granules by R. niveus glucoamylase also varied with the type of starch [105]. Reactions of glucoamylase on wheat and corn (maize) starch granules were followed visually by scanning electron microscopy and chemically by the determination of the amount of glucose released [106]. The scanning electron microscopy showed that wheat starch granules were attacked along the equatorial groove of the pancake‐shaped granules and that extensive reaction with maize starch granules produced a Swiss-cheese appearance with many deep holes into the granules [106].

A systematic study of the kinetics of glucoamylase was made using seven kinds of starch granules, and it was found that they divided into three groups: waxy maize starch, which was the most susceptible, being converted with 200 units into 98% glucose in 32 h; barley, maize, and tapioca starches were intermediate, being converted into 82, 77, and 75% glucose, respectively, in 32 h of reaction; and the third group of amylomaize‑7, shoti, and potato starches, which were the least susceptible, being converted into 21, 15, and 13%, respectively, in 32 h [107].

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Figure6

Scanning electron micrographs of maize starch granules that have been reacted with Aspergillus niger glucoamylase. A, 50 mg mL−1 starch granules, 20 IU mL−1 glucoamylase at 37 °C for 32 h; B, 50 mg mL−1 starch granules, 100 IU mL−1 glucoamylase at 37 °C for 32 h

Scanning electron microscopy showed that when the starch granules in the first two categories were converted into ∼50% glucose, there were numerous deep holes formed into the granules (see Fig. 6). Higher degrees of conversion showed that the remaining granules were hollow shells, with the interior parts hydrolyzed. It was concluded that glucoamylase enters the granule and catalyzed reactions inside the granule [106,107]. The granules of the least susceptible starches in the third group did not show much change in the granule morphology, except for some minor pitting of the surface.

When the starches in the third group from potato, shoti, and amylomaize‑7 were heated in water to their onset gelatinization temperatures of 60, 70, and 90 °C, respectively, for 0.5, 1, 2, and 4 h, the granules increased in size ∼2–3 fold and birefringence was lost. Glucoamylase hydrolysis of potato and amylomaize‑7 gelatinized starches was significantly increased after heating, from 13 to 86.6% for potato starch and from 21 to 45% for amylomaize‑7 starch. The glucoamylase hydrolysis of shoti starch, however, only increased slightly from 14.7 to 21.1%, after being heated for 4 h. The results after 4 h of heating of shoti starch granules were only slightly higher than the results after 0.5 h [108].

Two of the three starches that were quite resistant to glucoamylase hydrolysis, amylomaize‑7 and shoti starches, gave the highest percent reaction with the α‑(1→6) debranching enzyme, Ps. amyloderamosa isoamylase of 11.9 and 11.5%, respectively [109]. Waxy maize, maize, and barley starches were intermediate in their susceptibility to isoamylase hydrolysis, giving 7.8, 7.3, and 6.2%, respectively.

Heating at the gelatinization temperatures of the starch granules for 1 h significantly increased the reaction with isoamylase: potato starch increased from 3.7 to 56%; amylomaize‑7 starch increased from 11.9 to 36%; waxy maize starch was increased from 7.8 to 36%; and shoti starch was increased from 11.6 to 22%.

When gelatinized waxy maize starch was hydrolyzed 36% with isoamylase, only 16% of the hydrolyzed chains remained inside the granule. When 40% (v/v) ethanol was added to the waxy maize reaction digest, however, 44% of the isoamylase hydrolyzed chains remained inside the granule, and when 80% (v/v) ethanol was added, 100% of the hydrolyzed chains remained inside the granule [109]. With an 80% ethanol concentration, 100% of the isoamylase hydrolyzed chains also remained inside the granules for potato and shoti starches, and 98% of the isoamylase hydrolyzed chains remained inside the granules for amylomaize‑7 starch for a total percent isoamylase hydrolysis of 41, 10, and 13%, respectively [109].

An in situ method has also been developed to retain 100% of the glucose inside the granule by reacting it with glucoamylase in a minimal amount of water, 50 mg of starch in 1.0 mL of buffer, containing glucoamylase [110]. This was called the “solid-granule” reaction system. The buffer, with glucoamylase, completely enters the granules and reaction takes place inside the granules. Although eventually the concentration of glucose produced in the granules is relatively high, only a very low amount (< 0.01%) of reversion products (maltose, isomaltose, nigerose) were formed. Using a sealed vessel, it was possible to convert waxy maize starch granules into 51% glucose, all inside the granules. The amount of starch conversion was proportional to the number of non-reducing ends. It was possible to control the percent conversion by heating the reaction vessel at different times for 20 min at 110–120 °C to inactivate the glucoamylase and thereby control the amount of glucose produced to give 10, 20, 30, 40, and 50% hydrolysis. The granules with 20% or more glucose have a sweet taste [110].

A similar reaction was conducted with cyclomaltodextrin glucanyltransferase (CGTase) reacting with waxy maize starch granules in a minimal aqueous environment. The yield was disappointing, giving only 1.3% (w/w) cyclomaltodextrins inside the granules. This could be increased to 3.4%, with 100% retention of cyclomaltodextrin inside the granule by adding a combination of CGTase and isoamylase to the solid-granule reaction system [111].

In general, the rates of reaction of enzymes with native starch granules is 3- to 4‑orders of magnitude slower than they are with solubilized starch. The reaction of starch granules with enzymes is one method of modifying and altering the structure of starch granules. See Sect. 9 for chemical methods of modifying starch granules. It also is important to recognize that a single enzyme, reacting with starch granules from different plant sources or with solubilized starches from different plant sources will have significantly different activities on the different starches [26]. Furthermore, this activity can be greatly increased and the enzyme stabilized, by the addition of 0.04% (w/v) polyethylene glycol [26], as a specific-sized polyethylene glycol will give the maximum enzyme activity for a specific enzyme, however, the activities will still remain significantly different on the different starches [112].

35.8 Biosynthesis of Starch

Starch is biosynthesized in plant organelles. In leaf or transitory starch, the organelles are chloroplasts and in storage starch in seeds, tubers, stems, and roots, the organelles are amyloplasts. The process of starch biosynthesis is divided, for convenience of discussion, into three steps: (1) initiation, (2) elongation, and (3) termination, and for amylopectin, a fourth step, branching.

Starch chains were the first polysaccharides to have a mechanism proposed for their biosynthesis. This was in 1940 and was based on the studies of Hanes on the action of potato phosphorylase [113]. Hanes proposed that potato phosphorylase transferred glucose from α‑d‑glucopyranosyl‑1‑phosphate (α‑Glc‑1‑P) to the non-reducing ends of the starch chains and maltodextrins to give elongation of the starch chains. For starch biosynthesis by phosphorylase, it was recognized that the synthesis required a primer molecule that was either amylose, amylopectin, or a maltodextrin with a minimum of three glucose residues [114,115].

Phosphorylase is an enzyme that catalyzes a reaction that is reversible. The reaction can be either degradative or synthetic, depending on the starting substrates. It is degradative, if it is inorganic phosphate (Pi) or synthetic if it is α‑Glc‑1‑P, as shown in the following reaction:
$$ \text{P}_\text{i} + \underset{\text{starch chain}}{\text{G}-\text{G}-\text{G}-\text{G}-\text{G}-}\cdots \rlap{$\xrightarrow{\kern2mm\text{degradative}\kern1mm}$} \kern5.5mm\raisebox{-1ex}{$\xleftarrow[\text{synthetic}]{}$} \ \underset{\alpha\text{-Glc-1-P}}{\text{G}-\text{P}} + \ \underset{\text{degraded starch chain\hspace{-2.4mm}\llap{\raisebox{-2ex}{(putative primer)}}}}{\text{G}-\text{G}-\text{G}-\text{G}-\cdots}$$
(1)

The synthetic reaction of the putative primer chain, however, only will occur when the ratio of Pi to α‑Glc‑1‑P is less than the equilibrium value, which is 10.8 at pH 5, 6.7 at pH 6, and 3.1 at pH 7.0 [116]. These ratios are not obtained in vivo, as the concentration of Pi is 20- to 40‑fold higher than the concentration of α‑Glc‑1‑P [117,118] so that the in vivo conditions greatly favor degradation by phosphorylase, rather than synthesis.

Because a degraded starch chain is the product of the degradation reaction, the synthetic phosphorylase catalyzed reaction would require a starch chain or a maltodextrin chain to be a substrate in the synthetic reaction, as shown above. This was the origin for the primer required reaction for starch biosynthesis from the non-reducing-end of the primer, and it has pretty much been retained for 60 years [119,120,121].

In 1960, some 20 years after the phosphorylase experiments, it was found that UDP-Glc and ADPGlc were the high-energy glucosyl donors for the biosynthesis of starch chains, rather than α‑Glc‑1‑P, and that active starch synthesizing enzymes, starch synthase and starch branching enzyme were entrapped inside the starch granules [122,123]. When ADP‑[14C]Glc was incubated with starch granules, 14 C‑labeled glucose was incorporated into the starch in the granules. When this labeled starch was reacted with the exo-acting beta-amylase, 14 C‑labeled maltose was obtained and it was assumed that the glucose from ADPGlc was, therefore, being added to the non-reducing-ends of starch primers [122,123,124]. This experiment has been widely considered as proof that starch is biosynthesized by the addition of glucose from ADPGlc to the non-reducing-ends of starch primer chains. This conclusion, however, is not necessarily correct in that if starch chains were synthesized from the reducing-end de novo, independent of a primer, the synthesized starch chains would have every glucose residue labeled when the starch granules were reacted with ADP-[14C]Glc and reaction of this chain with beta-amylase would also give 14C-labeled maltose.

Because of these problems and the lack of definitive experiments supporting the mechanism of starch chain elongation, Mukerjea and Robyt carried out a number of experiments to test the non-reducing-end, primer mechanism [125,126]. The first series of experiments was the pulsing of starch granules from eight different plant sources with ADP‑[14C]Glc and the chasing of the pulsed starch granules with non-labeled ADPGlc [125]. The pulsed starch granules gave starch chains that on reduction with NaBH4 gave 14 C‑labeled d‑glucitol . The chased starch gave a significant decrease of 14 C‑labeled d‑glucitol. These experiments indicated that the starch chains were being elongated by the addition of glucose to the reducing-end and not to the non-reducing end, because if the glucose was being added to the non-reducing-end of a primer, it would be impossible to obtain any 14 C‑labeled d‑glucitol and further it would also be impossible to chase the label from the d‑glucitol. In addition, it was found that both labeled‐amylose and labeled‐amylopectin components were synthesized and that a significant amount of starch was synthesized: maize starch had an avg. labeled d.p. of 827, waxy maize 436, taro 462, rice 467, wheat 476, and potato 524. There was also evidence that covalent intermediates were formed with the starch synthase during synthesis [125].

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Figure7

Proposed mechanism for the elongation of starch chains by starch synthase, showing the initiation step (I) and the polymerization steps (II) in which d‑glucopyranose residues (circles) are added to the active site groups (X ) from ADPGlc and inserted between X and the reducing-end of a growing glucan chain by a transglycosylation reaction to give a polymer of α‑(1→4) linked d‑glucopyranose residues. The polymerization is terminated (III) when water enters the active-site and hydrolyzes the X‑acetal linkage of a reducing-end glucose residue attached to the starch chain. The mechanism gives a primer free elongation of the starch chain from the reducing-end. The dark circles are glucose residues and the dark circle with a line through it is a free hemi-acetal, reducing-end glucose residue

A two-site insertion mechanism for starch biosynthesis from the reducing-end was proposed, as shown in Fig. 7. The initiation step occurs with the glycosylation of the two sites, with the two nucleophilic X‑groups attacking C1 of the glucose residues of ADPGlc. Polymerization proceeds by the C4–OH group of one of the covalently linked glucose residues attacking the C1 group of the other, displacing the X‑group, which attacks C1 of another ADPGlc. The C4‑OH of this glucose residue then attacks the C1 of the maltosyl unit, which is transferred to the glucose residue, freeing the other X‑group, which makes an attack onto the glucose residue of another ADPGlc. This process continues, going back and forth between the two X‑groups, giving the polymerization of the starch chain by addition of glucose from ADPGlc to the reducing-end of the growing chain. The starch chain is extruded from the active site of starch synthase. The elongation of the chain is eventually terminated by hydrolysis of the chain from the active site.

In the second set of experiments by Mukerjea and Robyt, the effect of adding the putative primers to the reaction of ADP‑[14C]Glc was studied with three kinds of starch granules from maize, wheat, and rice [126]. The reactions were examined in the absence and presence of increasing concentrations of maltose (G2), maltotriose (G3), and maltodextrin (d.p. 12). All of the added putative primers were found to inhibit starch synthesis in increasing amounts, as the concentrations of the maltodextrins were increased, rather than stimulating synthesis, as would be expected for primers [126].

The putative primers did undergo a reaction. The major product in the presence of G2 was G3, with exponentially decreasing amounts of G4–G12; in the presence of G3, the major product was G4, with exponentially decreasing amounts of G5–G12. Further, the addition of glucose was to the non-reducing end of the putative primers. It was concluded that the putative primers were undergoing an alternate reaction in which they were acceptors that released glucose from the active site of starch synthase, with the glucose being added to C4–OH of their non-reducing-ends. This was confirmed by adding the acceptors to pulsed starch granules and observing the release of 14 C‑label from the granules in the absence of ADPGlc [126]. This study showed that primers are not involved in starch biosynthesis and in fact their presence inhibited starch biosynthesis.

Leloir and co-workers [122,123,124] and Frydman and Cardini [127] also found that the incubation of starch granules with maltodextrins and UDPGlc or ADPGlc exclusively transferred glucose to the non-reducing-ends of maltose, maltotriose, and maltotetraose primarily to give the next higher homologues, but they concluded that this was due to the primer non-reducing-end mechanism and did not recognize that starch synthesis was inhibited. This transfer of glucose to the non-reducing-end of carbohydrates is a well-known reaction in the synthesis of dextran by dextransucrase when low molecular carbohydrates are added to the reaction digest, glucose is transferred to C6–OH of the non-reducing-ends of the carbohydrates [128,129]. The carbohydrates are acceptors that inhibit dextran synthesis [129] just as the putative primers in starch biosynthesis accepted glucose and inhibited starch biosynthesis [126].

The third set of experiments by Mukerjea and Robyt involved two reactions with three varieties of starch granules from maize, wheat, and rice. In reaction I, the granules were reacted with 1‑mM ADP‑[14C]Glc and in reaction II, a portion of the granules from reaction I was reacted with 1‑mM nonlabeled ADPGlc [130]. The starch granules from the two reactions were solubilized and reacted with the exo-acting enzymes, glucoamylase and beta-amylase to an extent of 50% or less of the 14 C‑label. The amounts of 14 C‑labeled products from the two enzymes, respectively, were nearly equal for reaction I and reaction II. If the addition of glucose had been to the non-reducing-ends of primers, reaction II would not have given any labeled products. These results confirm that the elongation of the starch chain is the addition of glucose to the reducing-end by a de novo reaction and not by the addition of glucose to the non-reducing-end of a primer [130].

It must be emphasized that the mechanism of starch biosynthesis does not require a preformed starch chain primer or a maltodextrin chain primer and the starch chains begin de novo by forming covalent di-glucosyl intermediates from ADPGlc with starch synthase that proceed to synthesize amylose chains by adding glucose to the reducing-end of a growing chain, essentially by a transferase reaction of the enzyme covalently linked starch chain to the enzyme covalently linked glucose (see Fig. 7).

To produce amylopectin, it is necessary to have a starch branching enzyme that is capable of introducing the α‑(1→6) branch linkages into the linear amylose molecules synthesized by starch synthase. Such an enzyme (Q‑enzyme ) was first identified in potatoes [131] and then in broad beans [132]. The potato Q‑enzyme has been isolated and purified and the properties determined [133]. Q‑enzyme is a glucanyltransferase and cleaves an α‑(1→4) linkage of an amylose chain, and then transfers the chain to the C6–OH of another amylose-chain, rather than being transferred and attached to the residual part of the cleaved chain [134]. It was shown that water is not involved in these branching reactions [133].

There remain a number of important questions as to how branching enzymes convert the synthesized amylose chains into amylopectin and how the branch linkages appear in clusters in amylopectin, with a repeat distance of 70 Å. Another question is how starch granules are initiated and formed and how the percentages of amylose and amylopectin molecules are in a constant ratio that is specific for the particular plant source of the starch (see Table 2).

A logical assumption that can be surmised is that the enzymes that are found entrapped in the starch granules have most likely been responsible for the synthesis of the starch molecules in the granules. It has been hypothesized that starch synthase might exist both as a single enzyme and as a multi-enzyme complex of several starch synthases and branching enzymes. In the former case, amylose would be produced, and in the latter case, the multi-enzyme complex could produce amylopectin [135,136].

35.9 Chemical Modification of Starch

The chemical and physical properties of starch can be significantly altered by chemical modifications.

35.9.1 Acid Modifications of Starch Granules

One of the first reported modifications of starch granules was the treatment with aqueous acid in which there was hydrolysis of some or all of the glycosidic linkages. Obviously the molecular size of the polysaccharide molecules is decreased by acid hydrolysis. In 1874, Nägeli reported the treatment of native starch granules with 15% (w/v) sulfuric acid for one month at ∼20 °C. An acid‐resistant material resulted that was insoluble, but when isolated it could be solubilized in hot water [137]. This resistant material was called Nägeli Amylodextrin . It has an avg. d.p. of 24–30 and was fractionated by gel permeation chromatography to give three fractions [138]. Fraction I had the highest molecular weight and consisted of a mixture of branched maltodextrins with two or more branches per molecule. Fraction II, with a d.p. of ∼ 25, primarily had a single branch linkage, with the branch located close to the reducing end [138]. Fraction III was linear with an avg. d.p. of 13 [138]. With continued acid hydrolysis, Fraction I disappears and Fractions II and III are better resolved on Sephadex G‑50.

A soluble starch of much higher molecular weight than the Nägeli Amylodextrin was prepared by treating a suspension of starch granules with 7.5% (w/v) HCl for 1 week at 22–24 °C, or 3 days at 40 °C, followed by washing the granules free of acid [139]. The method is called the Lintner Procedure and is usually performed on potato starch granules. The Lintner Procedure is the method still used today to produce commercial soluble starch . The product is not completely water soluble and is polydisperse, with a relatively high reducing-value. The amylose component of Lintner soluble starch has an avg. d.p. of 340 and the amylopectin component has an avg. d.p. of 603, with a wide distribution of d.p. values [140].

Another method was reported in 1919 for obtaining soluble starch by refluxing the starch granules in 95% ethanol, containing 0.2–1.6% (w/v) HCl for 6–15 min [141]. The product is called Small's soluble starch . It is soluble (10 mg ml−1) in hot water and has a low reducing value due to the formation of ethyl‐glycosides at the hydrolytic points.

In 1987, Ma and Robyt reported the modification of starch granules by treating them with 0.36% (w/v) HCl in anhydrous methanol, ethanol, 2‑propanol, and 1‑butanol at 65 °C for 1 h. The starch granule was maintained, but the molecular sizes of the starch molecules progressively decreased in the different alcohols, as the number of carbons in the alcohols increased [140]. The molecular size distribution was greatly narrowed for these acid‐hydrolyzed starches and the amylose component completely disappeared in the starch granules modified in 2‑propanol and 1‑butanol. The starch granules modified in 1‑butanol also became quite soft. It was also found that acid hydrolysis was occurring inside the granules where there was 10–15% (w/w) water and the amount of acid also increased inside the granules as the size of the alcohols increased [140].

Fox and Robyt reported the kinetics of the acid-alcohol modifications by determining the avg. d.p. vs. time of reaction of the starch granules with 0.36 and 6.0% (w/v) HCl at 25 °C [142]. The kinetics showed that the avg. d.p. values dropped rapidly in the first 10 h and then leveled off, becoming constant at ∼30 h, d.p. values decreasing as the size of the alcohols increased and as the concentration of the acid increased. The avg. d.p. values varied from a high of 1717 to a low of 19, depending on the kind of alcohol and the acid concentration. The products represented new kinds of acid‐hydrolyzed limit dextrins, with a relatively narrow distribution and widely different avg. d.p. values, and different proportions of amylose and amylopectin.

The acid hydrolysis of starch granules in the four alcohols was also studied as a variation of temperature from 5 to 65 °C, acid concentration from 0.36 to 5.0% (w/v), and in the concentration of the starch granules [143]. The avg. d.p. values of the limit dextrins were dependent on all three parameters: first, the acid concentration and the temperature, of which combinations can be obtained to give limit dextrins with avg. d.p. values ranging from 1800 to 12 for the starch granules. In addition, the concentration of the starch granules was inversely proportional to the avg. d.p. values. This latter result confirmed that the mechanism of the hydrolysis of the glycosidic bond was taking place by the water inside the granules, because as the concentration of the starch granules increased, the concentration of the acid decreased inside the granules, due to the fact that there were more granules.

The acid hydrolysis of potato starch granules was studied using mixtures of two alcohols, methanol and 2‑propanol from 90:10 to 10:90 in 10% (v/v) intervals [144]. The avg. d.p. of the different modified starches leveled off in 48–72 h of reaction at 20 °C. The results gave a family of 11 curves, starting with 100% methanol and ending with 100% 2‑propanol, all with different limiting avg. d.p. values. As the concentration of 2‑propanol was increased and methanol decreased, the plots of avg. d.p. of the limit dextrins vs. the ratios of the two alcohols resembled titration curves, with different plateaus, in which the d.p. of the limit dextrin changed very slowly, or not at all. These plots were characteristic and dependent on the types of alcohols, the volume ratios of the two alcohols in the mixture, and the kind of starch [144]. It was proposed that the different ratios of the two alcohols produced the exposure of different glycosidic linkages to acid hydrolysis by converting different crystalline regions in the granule into amorphous regions that became susceptible to acid hydrolysis.

Another type of acid-modified starch is “thin-boiling” starches. Thin-boiling starches are produced by mild hydrolysis with 0.1–0.2% (w/v) HCl at 45–50 °C [145]. The treatment weakens the forces, holding the starch molecules together in the granule to give a starch that is more easily solubilized and has decreased viscosity. In the production of thin-boiling starches, the viscosity is monitored and the process is stopped when a particular desired viscosity is attained. The thin-boiling starches are widely used in the food industries to improve the appearance (clarity), stability, and texture of the food products in soups, sauces, dressings, bakery products, dairy products, and confectioneries.

35.9.2 Chemical Modification of the Starch Molecules in the Granules

The majority of the d‑glucopyranose residues in starch granules have three hydroxyl groups, one each at C‑2, C‑3, and C‑6, which can be modified. The amount of modification is called the “degree of substitution” (d.s.). If all three of the hydroxyl groups on all of the glucose residues are modified, the d.s. = 3; if two are modified, the d.s. = 2; and if only one is modified, the d.s. = 1; if one hydroxyl group out of every ten glucose residues is modified, the d.s. = 0.1, and so forth.

Three general kinds of modifications can be obtained: oxidation , esterification , and etherification to enhance hydrophilic character, to introduce hydrophobic character, to introduce positive or negative charges, to cross-link the starch chains, and to introduce color to the starch. The purposes for the modifications is varied, for example, to lower the gelatinization temperature, to decrease or prevent retrogradation, that is to decrease or prevent precipitation from solution, to enhance gelatinization, to increase the hydration of the starch molecules, to impart increased thickening and gelling properties, to increase binding of other molecules to starch, and to impart film-forming properties.

The function of modified starch is dependent on a number of things: the kind of starch (plant source), the amylose to amylopectin ratio, the nature of the substituents that are added, the degree of substitution, whether it is premodified by acid hydrolysis, and whether the starch is granular or gelatinized.

35.9.2.1 Oxidation of Starch

Alkaline hypochlorite is the most common method of oxidizing starch granules. The reaction is the oxidation of the secondary alcohols to ketones and the primary alcohol to a carboxyl group [146,147,148]. Alkaline hypochlorite oxidation is sometimes used to produce thin-boiling starches instead of acid hydrolysis.

Primary alcohol groups can be exclusively oxidized to aldehyde groups with pyridinium dichromate  [149,150] and to carboxyl groups with the 2,2,6,6‑tetramethyl‐1‑piperidine oxoammonium ion (TEMPO)  [151]. The aldehydes can then be reduced to primary alcohols by reaction with NaB3H4 [150,152], giving radiolabeled 3H‑starch and the carboxyl group can be inverted by the action of Azotobacter vinlandii poly‐β‑d‑mannuronic acid C‑5‑epimerase to give l‑iduronic acid  [153].

Vicinal alcohol groups, the 2,3‑glycol structure of starch, can be oxidized by periodic acid or periodate to cleave the carbon–carbon bond and oxidize the two carbons to produce two aldehyde groups at C‑2 and C‑3 [154,155]. The product is called “dialdehyde starch .” There are, however, very few actual aldehyde groups present. The main products are hydrated aldehyde groups and intra- and inter‐molecular hemiacetals [154,156,157]. The amount of oxidation can be controlled by the amount of periodate used. The resulting “aldehydes” can react with alcohols, amines, hydrazines, and hydrazides to give additional modification products. Chlorous acid will oxidize the “aldehyde groups” to polycarboxylic acid polymers [154].

35.9.2.2 Formation of Ester Derivatives

Common starch esters include the acetates. High d.s. starch acetates have been formed by using acetic anhydride with either sodium acetate or pyridine catalysts at 90–100 °C [158,159,160]. The major use of starch acetates have been in the study of the structures of amylose and amylopectin after acid hydrolysis. Acetylation of granular starch in aqueous suspension by acetic anhydride at pH 10–11 is used to produce low d.s. starch acetates that are primarily used for the stabilization of their viscosity and for their water-soluble clarity. The acetylation decreases the hydrophilic character and increases the hydrophobic character of starch. The major uses of starch acetates is in the paper industry for surface sizing to give improved print quality, uniform porosity, surface strength, and resistance to various solvents [161].

The solubility of starch acetates depends on the d.s. and the d.p. Starch acetates of 10–15% are soluble in water at 50–100 °C and are insoluble in organic solvents. Starch acetates of 40% or more are soluble in aromatic hydrocarbons and halogenated aliphatic hydrocarbons.

Starch esters formed by reaction of acid chlorides give a wider range in the kinds of esters that can be synthesized. In particular, reaction of oxalyl dichloride gives an ester with a free carboxylate group; reaction with succinyl dichloride also gives an ester with a free carboxylate group, but in addition can give intra- and inter‐molecular cross-linking of the starch chains; adipyl dichloride gives the same kind of reactions, but with greater potential for intermolecular cross-linking due to its larger size.

Starch granules have been cross-linked with phosphate by the reaction of an aqueous alkaline (pH 8–12) suspension reaction with phosphorus oxychloride [161]. Trimetaphosphate has also been used to produce phosphate cross-linkages. These reactions are primarily with the C‑6–OH groups and only a minimal number with the C‑3–OH groups [162]. Starch phospho esters can be obtained by phosphorylation with sodium triphosphate at pH of 8.5 [163].

The esterification of starch with p‑toleunesulfonyl (tosyl) chloride in pyridine gives partial 6‑O‑tosyl starch [164,165]. This ester can be replaced by a number of nucleophilic groups, such as iodide to give 6‑deoxy‐6‑iodo starch that in turn can be reduced with NaBH4 to give partial 6‑deoxy-starch or azide groups that on reduction give partial 6‑amino-6‑deoxy starch  [166,167].

35.9.2.3 Formation of Ether Derivatives

Ethers are formed with the alcohol groups of starch when alkyl halides or epoxides are added to an alkaline suspension of starch granules. The starch ethers that are produced are frequently water-soluble. Methyl or ethyl ethers can be formed by the reaction of starch at pH 10 with methyl or ethyl chlorides at 70–120 °C to give d.s. values of 1.2–2.3 [168]. A common ether is the formation of 6‑O‑carboxymethyl starch by the reaction of 2‑chloroacetic acid with alkaline starch [169]. Carboxymethyl starch is a water-soluble starch that is used as an emulsifier and thickening agent and provides stability against heat damage and amylase degradation.

Hydroxy ethers of starch are prepared by reaction of alkaline starch with epoxides, such as ethylene oxide and propylene oxide . The reaction is usually run on a 40–45% (w/v) suspension of starch granules in water under strong alkaline conditions (0.1–0.2‑M NaOH) at 40–50 °C, under nitrogen. The hydroxy ethyl starches are primarily used as binders for pigmented coatings and as surface sizing agents in paper manufacturing. Hydroxypropyl starches are of importance in food applications, where they are used as an edible, water-soluble film coating [161]. The formation of hydroxyalkyl derivatives increases their water solubility and decreases or prevents retrogradation of the starch chains. Ethylene oxide and propylene oxide do undergo “chaining reactions” in which they keep adding to the newly generated hydroxyl groups to give a chain of hydroxy alkyl ethers. This chaining reaction can be reduced or eliminated by conducting the reaction in the presence of methyl or ethyl chloride, which cap the newly formed hydroxyl groups, preventing further additions.

Starches can be chemically dyed by reacting a procion dye with 2,4,6‑trichloro‐triazine or 2,4‑dichloro‐triazine , followed by reaction with alkaline starch [170,171]. A chloro group on the triazine ring of the dye derivative is replaced by a starch hydroxyl group to form a stable ether linkage between the dye and the starch. Various colors of chlorotriazine dyes are available: Cibacron Blue, Procion Brilliant Blue, Procion Brilliant Red, Procion Brown, Procion Green, and Procion Yellow.

35.10 Analytical Methods in Starch Chemistry

35.10.1 Iodine/Iodide Colors

The formation of a blue-purple color by the interaction of starch and its components with iodine/iodide is a very characteristic and unique property of starch. The blue color that is observed is due to the amylose component and has a maximum absorbance at 645 nm. The other major component, amylopectin, gives a red-purple or maroon color, with a maximum absorbance at 490 nm. The ability of starch and its major components to give this iodine/iodide color provides a method for specifically detecting very low amounts of starch from as little as 1 µg mL−1. This property of starch is unique in that no other polysaccharide reacts with iodine/iodide to give a color and it hence provides a specific qualitative method for detecting starch and its components. It also provides a very sensitive method for detecting the cleavage of the α‑(1→4) glycosidic linkages produced by enzymatic and chemical degradations, as the color rapidly changes as the molecular size of the starch decreases [62].

The method: the iodine reagent to obtain the blue color of starch contains 0.2% (w/v) iodine in a 2.0% (w/v) potassium iodide solution [172]. Usually a solution with 10‑times this concentration is prepared and then diluted 1→10 immediately before use. The analysis is carried out by the addition of 25 µL of the diluted iodine/iodide reagent to 50 µL of each sample in a microtiter plate well and the absorbance is measured at 600–620 nm, using a microtiter plate colorimeter. The starch triiodide blue color is most stable under acidic conditions (pH 1–2) and will not form under alkaline conditions (pH 8 and higher).

It should be emphasized that the reaction to give the “blue color” is the reaction of starch with triiodide and higher complexes and not with iodine. The quantitative use of the starch‐triiodide color is problematic. As indicated, its most useful application is the qualitative identification of starch. An example is the identification of starch fractions from gel‐permeation chromatography [140,142] and the detection of the cleavage of the α‑(1→4) glycosidic linkages of starch by enzymatic or chemical degradation reactions, which gives a decrease and eventually a change in the color [65].

35.10.2 The Reducing Value

The reducing value , as the name implies, is the quantitative or qualitative measurement of the specific aldehyde or hemiacetal ends of the amylose and amylopectin molecules or the ends of their degradation products. There are several methods that might be used, such as the alkaline Somoygi–Nelson method  [173] or the ferricyanide/ferrocyanide/cyanide method  [174]. However, for amylose and amylopectin, there is only one reducing-end group per molecule, most of the reducing-value methods are not sensitive enough to determine the reducing-end of the starch macromolecules. In the last 15 years, however, a very sensitive, micro-method has been developed that can quantitatively measure the reducing-ends of the starch macromolecules when the starch is in concentrations of 1–9 mg mL−1 [175]. The method can also be used to measure the reducing value of smaller oligosaccharides, as well. It involves the copper‐bicinchoninate reagent as the oxidizing reagent that oxidizes the aldehyde and itself becomes reduced to give a purple color whose absorbance can be measured at 560 nm [175].

Two reagents are involved: Solution A consists of 97.1 mg of disodium 2,2′‑bicinchoninate dissolved in 45 mL of water containing 3.2 g of sodium carbonate monohydrate and 1.2 g of sodium bicarbonate, with the final volume made to 50 mL with water. Solution B consists of 62 mg of copper sulfate pentahydrate and 63 mg of l‑serine dissolved in 45 mL of water, with a final volume of 50 mL. The working reagent is prepared daily by mixing equal volumes of Solution A and B. Reducing carbohydrate standards consist of 1–20 µg mL−1.

The reducing value is determined by adding 100 µL of the reducing carbohydrate to 100 µL of the working reagent to the wells of a 96‑sample microtiter plate. The plate is covered with Saran wrap or some other method to seal the plate and prevent evaporation of the sample and reagent when the plate is heated in a water bath at 80 °C for 35 min. The plate is then cooled for 15 min and the absorbance is measured at 560 nm. Triplicate analyses are usually performed for each sample and for the maltose standards, along with a water blank [175].

It is important to use maltose or an equivalent disaccharide, such as isomaltose, cellobiose, nigerose, and so forth as standards, as glucose gives over‐oxidation for all of the reducing-value methods and cannot be used, unless glucose is the only carbohydrate reducing compound that is being measured. The substituted di- and higher‐oligosaccharides do not give over‐oxidation for most reducing value reagents, with the exception of the 3,5‑dinitrosalicylic acid method [176].

35.10.3 Determination of the Total Amount of Glucose-Carbohydrate in a Sample

The total amount of starch in a solution of pure starch can be measured, using the phenol‐sulfuric acid micro method  [175]. The procedure consists of adding 25 µL of sample (containing 10–200 µg of gluco‐carbohydrate mL−1) and 25 µL of 5% (w/v) phenol to the wells of a microtiter plate. Appropriate standards of glucose or maltose (10–200 µg mL−1) are used, along with a water blank. This should be done in triplicate. The plate is then placed into a bed of ice and 125 µL of concentrated sulfuric acid is added to each well, containing sample and phenol. The plate is again mixed at slow speed on a vortex mixer for 30 sec and then incubated in a water bath at 80 °C for 30 min and cooled, and the absorbance is measured at 492 nm.

35.10.4 Specific Determination of Starch When in the Presence of Other Carbohydrates, Using the Glucoamylase/Glucose Oxidase Method

The solution containing starch (∼1 g or less) in 100 mL of pyridinium acetate (pH 5.2) is hydrolyzed by 20 IU of glucoamylase (where IU equals the amount of enzyme that will produce 1 µmole of glucose at pH 5.2 and 37 °C in 1 min). The reaction is allowed to proceed at 37 °C for ∼5 h. These conditions should specifically hydrolyze 1 g of starch completely to glucose. The glucose is then specifically and quantitatively determined by the micro glucose oxidase method [175].

For the glucose oxidase determination of glucose, there are two solutions: Solution A consists of 7.5 mg of glucose oxidase and 0.75 mg of peroxidase (Sigma Chemical Co., St. Louis, MO, USA) which is dissolved in 12.5 mL of tris‐phosphate‐glycerol buffer (18.1 g of tris and 28.3 g of sodium dihydrogen phosphate dihydrate dissolved in 100 mL of water plus 100 mL of glycerol, diluted to 250 mL with water). Solution B consists of 50 mg of o‑dianisidine dissolved in 250 mL of the buffer described for Solution A above. The working reagent consists of equal volumes of Solutions A and B, prepared fresh daily. Solution A is stable for 1 week stored at 4 °C.

The analysis: 50 µL of sample containing 10–100 µg mL−1 is added to 200 µL of the working glucose oxidase reagent in a micro titer plate. Each sample is analyzed in triplicate and a blank is prepared with water. The plate is briefly mixed for 30 sec on a vortex plate mixer; 50 µL of concentrated HCl is added to each sample well and the absorbance is measured at 490–500 nm [175].

35.10.5 Determination of the Average Degree of Polymerization of Starch Fractions

The number of glucose residues in a starch component or fraction is termed the “degree of polymerization ” or d.p. The determination of the avg. d.p. is based on the counting of the number of molecules in a given weight of starch. A method of counting the number of molecules in a given sample is the reducing value as described in Sect. 10.2, using copper bicinchonate that “counts” the aldehyde/hemiacetal groups. The weight of the starch fraction can be obtained by using either the phenol‐sulfuric acid method (Sect. 10.3) or the glucoamylase/glucose oxidase method (Sect. 10.4). The avg. d.p. is calculated by the following formula:
$$ \makeatletter\ifx\letex\@undefined\def\textmu{$\mu$}\fi\makeatother\text{avg. d.p.} = \frac{\text{total weight of starch (\textmu g of glucose})}{\text{reducing value of starch (as \textmu g of maltose)}} \times 1.9 $$
(2)

The factor 1.9 is the quotient of the (molecular weight of maltose) / (molecular weight of glucose) or 342 / 180.

35.10.6 Determination of the Degree of Branching of Starch Fractions

A branched starch fraction whose degree of branching or percent of branching is to be determined, first has the avg. d.p. determined (Sect. 10.5). Then a sample of the branched compound is dissolved in 20‑mM acetate buffer (pH 4.8). Then 40 mIU of isoamylase per 50 µg (where 1 IU = 1 µmole of α‑(1→6) branch linkage is hydrolyzed per min by the enzyme). The isoamylase must be free of α‑amylase and glucosidase activities and can be obtained from Megazyme International [Wicklow, Ireland]. After reaction with isoamylase, the avg. d.p. is determined as per Sect. 10.5 The percent branching is then computed from the following:
$$ \text{\% branching} = \frac{\text{avg. d.p. after debranching with isoamylase}}{\text{avg. d.p. before debranching}} \times 100 $$
(3)
The average chain length of the branched chains in the starch fraction can then be calculated by the following:
$$ \text{avg. chain length} = \frac{1}{\text{\% branching}} \times 100 $$
(4)

If a sample has 5% branching, the avg. chain length would be (1 / 5) × 100 = 20 glucose residues. The individual maltodextrin chains can be separated and further analyzed by fluorescence‐assisted capillary electrophoresis (FACE), (see Sect. 15.3 in Chap. 1.2).

35.10.7 Measurement of Amylase Activity on Starch and Related Substrates

Amylases, both α‑ and β‑, hydrolyze the α‑(1→4) glycosidic linkages of starch to give reducing maltodextrin products that can be measured by using the micro copper‐bicinchoninate reducing value method (Sect. 10.2). Starch (1.5 mL of 0.5% w/v) is prepared in 25 mM of a buffer with the optimum pH of the α‑ or β‑amylase, containing 1.0 mM for α‑amylases; this is incubated for 10 min at the optimum temperature of the enzyme. The reaction is started by the addition of 25–50 µL of the enzyme to the starch solution. Aliquots of 150 µL are taken every 5 min for 30 min and added to 300 µL of 50 mM NaOH, which gives a pH of 12.5 and stops the reaction. The reducing value is measured, using the copper‐bicinchoninate method in which 100 µL of the stopped reaction is added to 100 µL of the working reagent in triplicate in a 96‑well microtiter plate. Similar amounts (100 µL) of maltose standards from 2.0–20.0 µg are added to 100 µL of the working reagent in triplicate. The plate is covered with Saran wrap to prevent evaporation and the plate is heated at 80 °C for 35 min, cooled, and the absorbance measured at 560 nm. Dilutions of the enzyme in the reaction mixture and the stopping reagent are used to determine the amount of reducing maltose produced by the α‑amylase and this is determined for each time point. The average of the triplicate measurements are obtained and a plot of µg of maltose produced versus the time of the reaction is prepared and the slope of the linear part of the line determined, which gives the µg of maltose produced per min. This is converted into µmoles of α‑(1→4) bonds hydrolyzed per min by dividing by the molecular weight of maltose (342), which gives the number of International Units of α‑ or β‑amylase [175].

35.10.8 Measurement of the Activity of Starch Synthase

Starch synthase catalyzes the synthesis of amylose. It can be measured in starch granules, where the enzyme is entrapped or it can be measured in a solution obtained in a plant extract.

35.10.8.1 Measurement of Starch Synthase Activity in Starch Granules

Starch granules (100 mg) are suspended in 1.0 mL of 0.1‑mM EDTA and 4‑mM glycine buffer (pH 8.4). The reaction is initiated by adding 20‑mM (0.2 µCi) ADP‑[14C]Glc and allowed to react for 1 h. The reaction is terminated by centrifuging and washing the granules 5‑times with 1 mL of water to obtain background radioactivity. This removes the unreacted ADP‑[14C]Glc from the granules and terminates the reaction. The granules are then treated 3‑times with 1 mL of anhydrous acetone to remove water. Small amounts of residual acetone are removed from the granules by pulling a vacuum for 1 min. The acetone-dried starch is weighed and the amount of 14 C‑glucose incorporated into the starch is determined by heterogeneous liquid scintillation counting by putting the granules in 10 mL of toluene scintillation cocktail [5.0 g PPO and 0.1 g POPOP in 1.0 L of toluene] and counting in a liquid scintillation spectrometer. The sample is counted for 10 min or 10,000 cpm, whichever comes first. The activity of the enzyme is nmoles of glucose incorporated into starch per h per 100 mg of starch granules [125].

35.10.8.2 Measurement of Soluble Starch Synthase Activity in a Plant Extract

Plant extract (25 µL) is added to 75 µL of 0.1‑mM EDTA and 4‑mM glycine buffer (pH 8.4), containing 20‑mM (0.05 µCi) ADP‑[14C]Glc. The digest is incubated at 37 °C for 30min. An aliquot (25 µL) is then added to (1.5 × 1.5 cm) Whatman 3‑mm paper and placed into 150 mL of methanol and stirred for 10 min. The washing is repeated two more times and the paper is then dried under a heat lamp. A blank is prepared with the 20‑mM (0.05 µCi) ADP‑[14C]Glc by adding 25 µL of water instead of plant extract and then putting 25 µL onto the Whatman 3‑mm paper and washing the paper 3‑times with 150 mL of methanol. The dried papers are added to 10 mL of toluene liquid scintillation cocktail and counted by heterogeneous liquid scintillation counting. The blank value is subtracted from the sample value to obtain the amount of 14 C‑glucose incorporated into starch.

35.10.9 Measurement of the Activity of Starch Branching Enzyme

The formation of amylopectin occurs through the action of starch branching enzyme on maltodextrins and/or amylose to form the α‑(1→6) branch linkage. Amylose or maltodextrins can be used as the substrates, but the amylose must be at a concentration of 1 mg/mL or less, as higher concentrations leads to the precipitation (retrogradation) of amylose from solution.

The assay described here uses amylose. Amylose (10 mg) is dissolved in 1.0‑mL dimethylsulfoxide by stirring on a hot plate and gently warming to 40–50 °C. After the amylose has been solubilized, the dimethylsulfoxide‐amylose solution is slowly diluted with 0.1‑mM EDTA and 4‑mM glycine buffer (pH 8.4) to 10.0 mL. The solution containing starch branching enzyme (100 µL) is added to 200 µL of the amylose solution and incubated at 37 °C for 30 min. The reaction digest is heated in a boiling water bath for 5 min, cooled, and 200 µL added to 200 µL isoamylase (250 mIU) and incubated 30 min at 37 °C. The isoamylase reaction digest (100 µL) is added in triplicate to 100 µL of the working reagent of the copper‐bicinchoninate and the reducing value determined, using maltose as a standard (see Sect. 10.2 for the reagent and the procedure). The activity of the starch branching enzyme is the number of α‑(1→6) bonds formed, which in this assay would be the µmoles of maltose determined min−1 × 1.9.

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© Springer-Verlag Berlin Heidelberg New York 2008
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