Analysis of nonstructural carbohydrates in storage organs of 30 ornamental geophytes by high-performance anion-exchange chromatography with pulsed amperometric detection


Author for correspondence:
W. B. Miller
Tel:+1 607 227 2780 Fax:+1 607 255 9998


  • • A comprehensive analysis of nonstructural carbohydrates in storage organs (bulbs and corms) of 30 ornamental geophytes was conducted by employing a variety of extraction techniques followed by high-performance anion-exchange chromatography with pulsed amperometric detection (HPAE-PAD).
  • • Among species, starch, fructan, glucomannan and soluble sugars accounted for 50–80% of storage organ dry weight (DW). Starch ranged from 24 to 760 mg g−1 DW, fructan (commonly occurring with starch) from 25 to 500 mg g−1 DW, and glucomannan from 15 to 145 mg g−1 DW. An acid hydrolysis protocol for concurrent determination of fructan and glucomannan was developed. The average degree of polymerization (DP) of ethanol and water-soluble fructan and the man : glu ratio of glucomannan also varied between species.
  • • The 80% ethanol fraction contained soluble sugars and short-chain fructans (< 25 DP), whereas water extracts contained soluble sugars, fructans (both short- and long-chain, ≤ 100 DP), and glucomannan. A substantial portion of the starch became ‘soluble’ in water during extraction, and depended on the species and extraction temperature.
  • • Our results indicate that extraction and analysis techniques of nonstructural carbohydrates for physiological and biochemical research on geophytic storage organs should be validated to accurately understand the identity of diverse carbohydrate pools, their physiological relevance and functions.


Geophytes have evolved to survive adverse environmental conditions through the production of underground storage organs that may also serve as a means of vegetative propagation. Storage of large amounts of reserve carbohydrate in the underground organs is critical for the geophytic growth strategy to ensure a supply of carbon and energy for maintenance during unfavorable conditions, and for rapid shoot and reproductive growth when environmental conditions are favorable. Starch is the most abundant reserve carbohydrate in geophytes; however, other reserve carbohydrates (e.g. fructan and glucomannan) also occur in place of, or in addition to, starch (Miller, 1992). In spite of the importance of storage carbohydrate metabolism in ornamental geophytes, and their worldwide economic significance, there is a paucity of information concerning their identity and distribution among species.

Fructans (linear or branched polymers of fructose with a terminal glucose unit) are the most common alternative to starch as a reserve carbohydrate in plants. It has been estimated that fructans occur as the principal reserve carbohydrate in 15% of contemporary angiosperm plant species (Hendry, 1993). These fructan-containing species can be found within a diverse range of families of both monocots and dicots, including Poaceae, Amaryllidaceae, Iridaceae, Liliaceae, Asteraeae and Campanulaceae (Hendry & Wallace, 1993). Most of these species have economic significance such as cereals (e.g. barley, oat and wheat), forage grasses (e.g. Lolium and Festuca), vegetables (e.g. onion, chicory and asparagus), and ornamentals (e.g. tulip, dahlia and hyacinth). In plant cells, fructans are stored in vacuoles, and it is assumed that they are synthesized there as well (Vijn & Smeekens, 1999). In addition to its role as a reserve carbohydrate, many other functions have been proposed for fructans, including osmoregulation, cryoprotection and sink regulation (Pollock, 1986; Hendry, 1987). The occurrence of fructans (especially in storage organs) is widespread in ornamental geophytes, indicating their importance in this group of plants (Hendry & Wallace, 1993). However, little information is available on the structure and biochemistry of fructans in this group of plants (Miller, 1992).

Glucomannans are either linear or branched heteropolymers of glucose and mannose that usually form gel-like mucilages. Glucomannans also occur as a reserve carbohydrate in several geophytes (Miller, 1992), although less frequently than fructans. For example, glucomannan has been identified in storage tissues of Lilium (Tomoda et al., 1976), Narcissus (Kato et al., 1973; Tomoda et al., 1980), and Amaryllis (Tomoda et al., 1985). Most of our knowledge on glucomannan comes from work on seed mannan (Bewley & Reid, 1985). The colloidal nature of glucomannans suggests a possible role in cellular water relations (Meier & Reid, 1982); however, no conclusive evidence on their specific functions in geophytes is available.

Development of techniques for accurate quantification and characterization of these different carbohydrates is needed for the advancement of research on their metabolism and function in geophytes. Specific HPLC methods have been developed to separate and detect fructan polymers, and gas-liquid chromatography-mass spectrometry (GLC-MS) has been used to characterize their linkage structures (Bancal et al., 1993). High-performance anion-exchange chromatography (HPAE) with pellicular resins performed under alkaline conditions, coupled with pulsed amperometric detection (PAD) on a gold working electrode, is a powerful technique for analyzing soluble carbohydrate polymers (Lee, 1990). A high degree of resolution and highly sensitive detection of soluble carbohydrates, including polymers, can be attained with HPAE-PAD without sample derivatization. HPAE-PAD has been a key tool in structural studies of plant fructan (Chatterton et al., 1990; Ernst et al., 1996; Pavis et al., 2001).

The solubility characteristics of starch, fructan and glucomannan are significantly different, and therefore the concurrent existence of these carbohydrates in geophytic storage organs poses a challenge for designing extraction protocols for accurate quantification of each carbohydrate fraction. A review of studies on reserve carbohydrates in geophytes reveals that this aspect has not been given careful attention. Commonly used protocols include extraction of soluble carbohydrates with hot ethanol (70–80%) followed by water extraction, or direct water extraction, and then determination of starch remaining in the residue. Mono- and disaccharides and only short-chain (low degree of polymerization, DP) fructans are usually soluble in ethanol, whereas most of the fructans (including higher-DP polymers) are soluble in water. Glucomannans are usually soluble in water but not in ethanol. Our preliminary experiments with several geophyte storage organs revealed the presence of significant amounts of soluble starch in water extracts. The presence of soluble starch (glucan polymers) in extracts interferes with the accurate quantification of fructan and glucomannan, and also leads to an underestimation of the starch content determined in the residue.

The objective of the present study was to perform a comprehensive qualitative and quantitative analysis of nonstructural carbohydrates present in storage organs of 30 different geophytes. Different extraction methods were compared to investigate the solubility of each carbohydrate fraction (soluble sugar, fructan, glucomannan and starch) and their effects on quantification techniques. HPAE-PAD was used to resolve and detect soluble carbohydrates, and chromatographic profiles of fructans of each species were compared.

Materials and Methods

Plant materials

Bulbs or corms of 30 species or cultivars (Table 1) of ornamental geophytes were obtained from a commercial source in the Netherlands in the autumn following normal commercial harvesting and transportation procedures. Upon receipt, all plant materials were stored at 17–20°C before use. Outer dry scales or tunics of bulbs were removed and scales were cleaned well with paper towels. Each bulb (or corm) was cut and the central bud was removed. Storage tissues (minus the central bud) were immediately frozen in liquid nitrogen and stored at −80°C until freeze-drying. Freeze-dried tissues were ground to a powder and stored at 20°C with desiccant.

Table 1.  List of geophytes used in the study
Species/cultivarFamilyStorage organ
Allium caeruleumAlliaceaeBulb
Allium christophiiAlliaceaeBulb
Allium hollandicum ‘Purple Sensation’AlliaceaeBulb
Allium karataviense ‘Ivory Queen’AlliaceaeBulb
Allium neapolitanumAlliaceaeBulb
Allium sphaerocephalonAlliaceaeBulb
Arum italicumAraceaeBulb
Cammassia leichtlinii ‘Caerulea’LiliaceaeBulb
Chionodoxa forbesiiLiliaceaeBulb
Colchicum autumnale ‘The Giant’LiliaceaeCorm
Crocus vernus ‘Pickwick’IridaceaeCorm
Fritillaria imperialis ‘Rubra Maxima’LiliaceaeBulb
Fritillaria persicaLiliaceaeBulb
Galanthus nivalisAmaryllidaceaeBulb
Hyacinthus orientalis ‘Carnegie’HyacinthaceaeBulb
Hyacinthus orientalis ‘Pink Pearl’HyacinthaceaeBulb
Iris bucharicaIridaceaeBulb
Iris reticulata ‘J.S. Dyt’IridaceaeBulb
Iris reticulata ‘Pauline’IridaceaeBulb
Iris xiphium ‘Blue Diamond’IridaceaeBulb
Muscari armeniacumLiliaceaeBulb
Narcissus ‘Carlton’AmaryllidaceaeBulb
Narcissus ‘ce Follies’AmaryllidaceaeBulb
Narcissus ‘Minnow’AmaryllidaceaeBulb
Narcissus ‘Tete a Tete’AmaryllidaceaeBulb
Scilla sibericaLiliaceaeBulb
Tulipa gesneriana ‘Apeldoorn’LiliaceaeBulb
Tulipa gesneriana ‘Monte Carlo’LiliaceaeBulb
Tulipa tardaLiliaceaeBulb
Tulipa turkestanicaLiliaceaeBulb

Extraction of nonstructural carbohydrates

Extraction with ethanol Tissue samples (50 mg dry tissue) were extracted at 70°C with 80% ethanol (three extractions of 3 ml each, 30 min per extraction). Tissue samples were weighed into conical centrifuge tubes and the extractant was added. Each tube was vortexed for 5 s and placed in a water bath. No sonication or shaking was done during the 30 min of extraction. At the end of each extraction, each tube was vortexed again for 5 s before centrifugation. Tissue suspensions were centrifuged at 4000 g for 10 min after each extraction, and the supernatants were combined. The extracts were passed through ion exchange columns consisting of 1 ml each of Amberlite IRA-67 (acetate form) (Sigma, St Louis, MO, USA) and Dowex 50W (hydrogen form) (Sigma) to remove charged material. The extracts were evaporated to dryness at 55°C under vacuum, and dissolved in 14 ml of HPLC-grade water. This fraction was referred as the ethanol-soluble fraction.

Extraction with water Tissue samples (50 mg dry tissue) were boiled for 5 min with HPLC-grade water (adjusted to pH 8.0 with CaCO3) and then extracted at 70°C (three extractions of 3 ml each, 30 min per extraction). Tissue suspensions were centrifuged at 4000 g for 10 min after each extraction, and the supernatants combined. The extracts were passed through ion exchange columns (as already described), and made up to 14 ml with HPLC-grade water. This fraction was referred as the water-soluble fraction.

Extraction with water after ethanol extraction  The residue after ethanol extraction was dried in an oven at 55°C overnight. The residue was extracted twice at 70°C with 5 ml of HPLC-grade water (adjusted to pH 8.0 with CaCO3) for 30 min each time. Supernatants were collected after centrifugation, pooled, and made up to 14 ml with water. This fraction was referred to as the water-soluble fraction after ethanol extraction.

Internal standard  When used, mannitol was added (1 mg per sample, at the start of the first extraction) as an internal standard for HPLC. Under normal HPLC conditions, it was cleanly resolved from any plant-derived peaks of interest. The internal standard was mainly useful with the initial alcohol extract when there is a possibility of sample loss in the procedure. In most cases, an internal standard was not used in the sequential extraction experiments, or in hydrolysis procedures.

Determination of starch  The residue remaining after ethanol or water extraction was boiled for 30 min with 4 ml of 100 mm Na-acetate buffer (pH 4.5) to gelatinize starch. After cooling, 50 units of amyloglucosidase (in 1 ml of Na-acetate buffer, pH 4.5; Sigma) were added to each sample and incubated for 24 h at 55°C. The amount of glucose released was determined by HPAE-PAD using an aliquot of the digested sample. The amount of starch was estimated according to the amount of glucose released. Experiments confirmed (data not presented) that the enzyme preparation used had no activity against citrus pectin.

Insoluble fructan or glucomannan  To determine the presence of any fructan or mannan that was not soluble in either ethanol or water, an aliquot of the residue suspension after ethanol and/or water extraction was directly digested with HCl under the conditions to be described. The resultant monosaccharides were determined with HPAE-PAD.

Acid hydrolysis of the extracts

Acid hydrolysis of the extracts was performed to determine the total amount of fructan and glucomannan, and also to determine the glucose : fructose ratio in fructan, and mannose : glucose ratio in glucomannan. In preliminary tests to determine optimum acid concentration and duration for hydrolysis of each fraction, samples were boiled with 0.05 m HCl for 5 min, or with 0.05 m, 0.2 m, 1.0 m or 2.0 m HCl for 30 min, or with 1.0 m or 2.0 m HCl for 4 h. Samples were cooled after hydrolysis and neutralized with NaOH. After appropriate dilution of the neutralized samples, an aliquot was subjected to HPAE-PAD to determine the resulting individual monosaccharide concentrations, and to check for the presence of any undigested peaks. According to the results of these preliminary tests, samples were hydrolyzed for 5 min with 0.05 m HCl for fructan determination, and for 30 min with 2.0 m HCl for glucomannan determination in routine experiments.

Starch solubility in water

Since preliminary experiments indicated that starch was soluble to different degrees in water depending on the species, a detailed experiment was conducted to determine the conditions under which starch became solubilized in water during extractions. Tissues (50 mg of dry tissue) were extracted with distilled water (pH adjusted to 8.0) at 25°C after 5 min of boiling, 25°C with no boiling, 70°C after 5 min of boiling, or 70°C after ethanol extraction. After 5 min of boiling, samples were brought to 25°C by immediately immersing the tubes in a 25°C water bath. In all cases, tissues were extracted twice for 30 min each at given temperatures, and suspensions were centrifuged at 4000 g for 10 min after each extraction to remove the supernatant. The starch remaining in the residue after extraction was determined after digestion with amyloglucosidase. The starch determined in the residue after the extraction with 80% ethanol served as the reference total starch for the calculation of percentage of starch soluble under each extraction method.


Carbohydrates were subjected to HPAE-PAD to resolve and quantify soluble sugars and oligosaccharides. A Dionex DX-500 series chromatography system, consisting of a GP50 gradient pump, an ED40 electrochemical detector with a gold working electrode, and an AS40 autosampler, was used. System control, data acquisition and processing were done with PeakNet (5.1) software (Dionex, Sunnyvale, CA, USA). A Carbopac PA-1 analytical (4 × 250 mm) column with a guard (4 × 50 mm) column (Dionex) was used for carbohydrate separation. Carbohydrates were eluted at a flow rate of 1.0 ml min−1 at c. 1600 psi. The electrode pulse potentials and durations of the waveform were as follows: E1 = 0.1 V, 400 ms; E2 = −2.0 V, 20 ms; E3 = 0.6 V, 10 ms; and E4 = −0.1, 70 ms. The following elution program was used to resolve glucose, fructose, sucrose and fructan polymers: 200 mm NaOH (0–5 min); a linear gradient from 0 to 500 mm of sodium acetate in 200 mm NaOH (5–25 min); 500 mm sodium acetate in 200 mm NaOH (25–28 min); a linear decreasing gradient from 500 to 0 mm of sodium acetate in 200 mm NaOH (28–29 min); 200 mm NaOH (29–32 min). Samples containing mannose were eluted with 18 mm NaOH for 25 min, followed by 15 min of column wash with 200 mm NaOH and 20 min column equilibration with 18 mm NaOH. These conditions were necessary to resolve glucose and mannose. The amounts of glucose, fructose, mannose and sucrose were determined by comparison with calibration curves derived from standard authentic sugars (Sigma). Since standards were not available for fructan polymers, quantification of each individual fructan polymer was not possible.

Calculations of different carbohydrates

The amounts of free glucose, fructose, mannose and sucrose in the samples were obtained directly from chromatograms before acid hydrolysis. For total fructans, the sum of fructose and glucose, resulting from acid hydrolysis, was calculated after subtracting the amounts of respective free monosaccharides (before hydrolysis) and monosaccharides originating from sucrose. For the calculation of total glucomannan, the mannose resulting from hydrolysis (after subtracting free mannose before hydrolysis) was added to the glucose resulting from hydrolysis after subtracting free glucose (before hydrolysis), and glucose originating from sucrose and fructan (if present). For the calculation of starch, the amount of glucose resulting from amyloglucosidase digestion was used. The monosaccharide totals were corrected for water molecules added during the hydrolysis to estimate the amounts of fructan, glucomannan and starch.


Extraction methods

In storage organs of all species, ethanol-soluble, water-soluble and water-soluble fraction after ethanol extraction were compared for the presence of different carbohydrates. A summary of the solubility characteristics is given in Table 2. The ethanol-soluble fraction contained only soluble sugars (mono- and disaccharides) and short-chain fructans (< c. 25 DP). Digestion with amyloglucosidase and acid hydrolysis with 2.0 µ HCl confirmed the absence of soluble starch and glucomannan, respectively, in ethanol extracts. Water extracts contained soluble sugars, fructans (both short- and long-chain, up to c. 100 DP), glucomannan and a varying quantity of starch (glucan polymers) depending on the species. The presence of glucans interfered with the acid digestion procedures for the determination of fructans and glucomannan. Extraction with water after ethanol extraction, however, substantially reduced the solubility of starch, while it completely solubilized long-chain fructans and glucomannan.

Table 2.  Summary of solubility characteristics of different types of nonstructural carbohydrates with different extraction methods
Type of carbohydrate80% ethanol extractionWater extractionWater extraction after 80% ethanol
Mono- and disaccharidesCompletely solubleCompletely solubleNot detected (all are solubilized in 80% ethanol)
FructanLow-DP fructans are soluble (< c. 25 DP)Both low- and high-DP-soluble (up to c.100 DP)High-DP fructans are soluble (c. 25–100 DP)
StarchNot solubleSolubility in range 0–97% depending on the species and temperatureSoluble to a lesser degree (> 20%)
GlucomannanNot solubleSolubleSoluble

More than 98% of total fructan and glucomannan in all species were solubilized by direct extraction with water (i.e. the amount of monosaccharides released by mild acid hydrolysis from tissue residue after water extraction accounted for < 2% of total ). The yields of extracted soluble sugars, total fructans and glucomannans obtained with direct water extraction were comparable to the sum of yields from ethanol extraction and water extraction after ethanol extraction in all the species, except in Galanthus nivalis (in the case of fructans) and Camassia leichtlinii (in the case of glucomannan). In G. nivalis, extraction with ethanol made 76% of the fructan insoluble (which would have been soluble if extracted with water directly), and in C. leichtlinii, extraction with ethanol made 13% of glucomannan insoluble during subsequent water extraction. Therefore, for those two species, direct water extraction is the best method to completely solubilize fructan and glucomannan.

Based on these findings, the following procedure was used for the final determination of concentrations of each nonstructural carbohydrate component. Tissues were first extracted with 80% hot ethanol followed by extraction with water. Soluble sugars (glucose, fructose and sucrose) were determined in the ethanol extract. Fructan was determined in both the ethanol extract, the water extract after ethanol extraction, and the tissue residue after extraction (by mild acid hydrolysis) and combined to calculate the total amount. Glucomannan was determined in the water extract obtained after ethanol extraction. In G. nivalis and C. leichtlinii, the direct water extract was used to determine soluble fructan and glucomannan concentrations, respectively. For starch, a separate set of samples was extracted with ethanol and the residue was used directly for digestion with amyloglucosidase. The amyloglucosidase preparations did not hydrolyze fructans, as evidenced by the absence of fructose peaks in HPAE-PAD chromatograms of digested fructan samples.

Acid hydrolysis of carbohydrates

Extracts containing different combinations of carbohydrates were selected (i.e. fructan only, fructan and glucomannan, and fructan and soluble starch) and hydrolyzed with different acid concentrations for various durations to determine optimum acid hydrolysis conditions for each type of carbohydrate. For example, Fig. 1 illustrates the resulting monosaccharide profiles when the water extract after ethanol extraction from C. leichtlinii was hydrolyzed with HCl under various conditions. The extract contained fructan and glucomannan only. Hydrolysis with 0.05 m HCl at 100°C resulted in the appearance of fructose and a small amount of glucose indicating the hydrolysis of fructan. With stronger hydrolysis conditions, the destruction of fructose was evident. Mannose appeared with 0.2 m HCl digestion for 30 min, then increased in concentration with stronger hydrolysis conditions, reaching a maximum with hydrolysis with 2.0 m HCl for 30 min. Similar acid hydrolysis patterns for fructan were observed with extracts from Hyacinthus orientalis (see Supporting Information, Fig. S1), Allium caeruleum and pure inulin from chicory root (Sigma; data not shown). Also, similar acid hydrolysis patterns (release of mannose) were observed with glucomannan extracts from Narcissus ‘Carlton’ and pure mannan from Saccharomyces cerevisiae (Sigma; data not shown). Water-soluble starch was also hydrolyzed by HCl. Glucose appeared with hydrolysis in 0.2 m HCl for 30 min in extracts containing soluble starch, and increased with stronger conditions (data not shown). A similar hydrolysis pattern was observed with potato-soluble starch (Sigma). This observation shows that the presence of soluble starch may interfere with acid hydrolysis treatments to determine glucomannan and would lead to overestimation of glucomannan content. According to these results, 0.05 m HCl for 5 min at 100°C was selected as the fructan hydrolysis treatment, and 2.0 m HCl for 30 min at 100°C was selected as the glucomannan hydrolysis treatment for final determinations. Known amounts of free fructose, glucose and mannose were digested under the conditions described to confirm that free monosaccharides were not destroyed by them (fructose with 0.05 m HCl for 5 min, and glucose and mannose with 2.0 m HCl for 30 min).

Figure 1.

Concentrations of hydrolysis products (monosaccharides) of water extract after ethanol extraction from Cammassia leichtlinii. Samples were hydrolyzed with HCl (at 100°C) under given conditions, and the resultant monosaccharide concentrations were determined by high-performance anion-exchange chromatography with pulsed amperometric detection (HPAE-PAD). Fructose, black bars; glucose, gray bars; mannose, white bars. Each bar is a mean ± SE, n = 3.

In all the fructan-containing samples, no significant differences were observed in the fructose yield when compared between hydrolysis with 0.05 m HCl for 5 min and that with 0.2 m HCl for 30 min (data not shown). Similar fructose yields were obtained when samples were digested with inulinase (fructozyme L) from Aspergillus niger (Sigma). The enzyme preparation, however, was not specific for fructan, as it hydrolyzed sucrose and potato-soluble starch to a considerable extent and pure mannan from Saccharomyces cerevisiae slightly (data not shown).

Nonstructural carbohydrates in geophytes

Starch and fructan were the major nonstructural carbohydrates detected in the geophytes investigated. Soluble sugars (glucose, fructose and sucrose) were present in all geophytes at concentrations < 100 mg g−1 DW (Table 3). Free mannose was detected only in C. leichtlinii (c. 0.2 mg g−1 DW). Among all species, the concentration of sucrose was always greater than that of glucose and fructose. Starch was present in all taxa except Allium sp. and C. leichtlinii. The concentration of starch varied from 244 mg g−1 DW (Muscari armeniacum) to 760 mg g−1 DW (Tulipa turkestanica). Fructan was present in 25 out of 30 geophytes investigated, and concentrations ranged from 23 to 508 mg g−1 DW. Allium species contained the highest concentration of fructan (about half of the DW). Fructan occurred concurrently with starch storage organs of 18 out of 30 geophytes.

Table 3.  Concentrations of various nonstructural carbohydrates in storage tissues of different geophyte species
Species/cultivarConcentration (mg g−1 DW)
  1. Values are mean ± SD, n = 3.

  2. nd, not detected.

Allium ‘Ivory Queen’0.6 ± 0.14.7 ± 0.418 ± 1nd439 ± 22nd
Allium ‘Purple Sensation’0.7 ± 0.12.8 ± 0.310 ± 1nd478 ± 16nd
Allium caeruleum0.4 ± 0.13.6 ± 0.518 ± 1nd458 ± 18nd
Allium christophii0.8 ± 0.14.2 ± 0.914 ± 1nd502 ± 38nd
Allium neapolitanum0.3 ± 0.11.0 ± 0.18 ± 1nd508 ± 16nd
Allium sphaerocephalon0.8 ± 0.17.4 ± 1.020 ± 1nd451 ± 15nd
Arum italicum1.3 ± 0.11.2 ± 0.131 ± 2699 ± 19nd19 ± 1
Camassia forbesii0.8 ± 0.11.3 ± 0.129 ± 2539 ± 184 ± 4nd
Camassia leichtlinii ‘Caerulea’1.2 ± 0.16.2 ± 0.241 ± 2nd353 ± 9146 ± 6
Colchicum ‘The Giant’2.9 ± 0.22.6 ± 0.282 ± 6608 ± 1ndnd
Crocus vernus ‘Pickwick’6.4 ± 0.33.3 ± 0.222 ± 1667 ± 16 ndnd
Fritillaria imperialis ‘Rubra Maxima’1.5 ± 0.52.2 ± 0.429 ± 1724 ± 8ndnd
Fritillaria persica3.3 ± 0.54.2 ± 0.842 ± 2677 ± 28ndnd
Galanthus nivalis0.6 ± 0.11.4 ± 0.127 ± 1492 ± 25149 ± 10nd
Hyacinthus orientalis ‘Carnegie’0.7 ± 0.14.3 ± 0.219 ± 1356 ± 17286 ± 1316 ± 1
Hyacinthus orientalis ‘Pink Pearl’0.7 ± 0.14.6 ± 0.122 ± 2355 ± 17284 ± 515 ± 1
Iris bucharica2.9 ± 0.22.0 ± 0.222 ± 1471 ± 252 ± 3nd
Iris reticulata ‘J.S. Dyt’0.7 ± 0.13.8 ± 0.624 ± 1539 ± 25157 ± 13nd
Iris reticulata ‘Pauline’1.1 ± 0.17.6 ± 1.423 ± 4534 ± 22152 ± 5nd
Iris xiphium ‘Blue Diamond’2.3 ± 0.38.3 ± 0.680 ± 4485 ± 2107 ± 9nd
Muscari armeniacum0.6 ± 0.14.4 ± 0.416 ± 2244 ± 1362 ± 21nd
Narcissus ‘Carlton’1.5 ± 0.12.6 ± 0.129 ± 1685 ± 1033 ± 141 ± 3
Narcissus ‘Ice Follies’1.4 ± 0.12.3 ± 0.227 ± 1656 ± 2929 ± 436 ± 1
Narcissus ‘Minnow’1.0 ± 0.13.3 ± 0.329 ± 1564 ± 2285 ± 2 32 ± 2
Narcissus ‘Tete a Tete’0.9 ± 0.11.9 ± 0.330 ± 1664 ± 1530 ± 14 ± 1
Scilla siberica0.9 ± 0.12.5 ± 0.729 ± 2473 ± 2184 ± 1423 ± 1
Tulipa gesneriana ‘Apeldoorn’9.3 ± 0.74.5 ± 0.156 ± 2687 ± 1954 ± 5nd
Tulipa gesneriana ‘Monte Carlo’2.5 ± 0.21.7 ± 0.144 ± 1711 ± 1744 ± 3nd
Tulipa tarda2.1 ± 0.32.9 ± 0.348 ± 3711 ± 3533 ± 4nd
Tulipa turkestanica0.8 ± 0.11.1 ± 0.132 ± 2760 ± 3123 ± 2nd

The solubility and the polymer size of fructan varied considerably among the geophytes (Table 4). Assuming that each fructan molecule contains one glucose unit, the fructose : glucose ratio provides an estimate for the average DP of fructan. Tulipa and Narcissus mainly contained short-chain fructan, of which > 95% was soluble in ethanol. The average DP of ethanol-soluble fructan ranged from 3.6 to 34.4 (usually < 20 DP), except in Muscari armeniacum and Scilla siberica, where the average DP of water-soluble fructan varied from 23 to 109. Interestingly, the ratio of ethanol-soluble to water-soluble fructan varied considerably among the Allium species.

Table 4.  Solubility characteristics and fructose : glucose ratio of fructans in different geophyte species
Species/cultivar% of fructanFructose : glucose ratio
Soluble in 80% ethanolSoluble in water after ethanol extractionRemaining in tissue after extractionEthanol-soluble fructanWater-soluble fructan
  1. Values are means, n = 3.

  2. nd, could not be determined accurately because of the low amount of fructan present.

Allium ‘Ivory Queen’
Allium ‘Purple Sensation’29.469.31.312.585.9
Allium caeruleum58.740.9 0.437.8
Allium christophii41.157.61.313.695.8
Allium neapolitanum8.289.91.910.8108.6
Allium sphaerocephalon79.420.30.314.438.3
Camassia forbesii35.563.11.410.759.9
Camassia leichtlinii ‘Caerulea’
Galanthus. nivalis6.816.876.45.432.6
Hyacinthus orientalis ‘Carnegie’
Hyacinthus orientalis ‘Pink Pearl’43.655.80.615.559.9
Iris bucharica64.831.33.94.823.6
Iris reticulata ‘J.S. Dyt’
Iris reticulata ‘Pauline’
Iris xiphium ‘Blue Diamond’
Muscari armeniacum41.655.62.721.479.2
Narcissus ‘Carlton’
Narcissus ‘Ice Follies’
Narcissus ‘Minnow’
Narcissus ‘Tete a Tete’
Scilla siberica39.459.11.534.492.8
Tulipa gesneriana ‘Apeldoorn’
Tulipa gesneriana ‘Monte Carlo’
Tulipa tarda99.
Tulipa turkestanica98.

Glucomannan was present in nine geophytes, including H. orientalis, Narcissus sp., Arum italicum, C. leichtlinii and S. siberica. The concentrations were, however, far lower than those of starch or fructan, with the highest concentration in C. leichtlinii (146 mg g−1 DW). More than 99% of glucomannan was soluble in water except in C. leichtlinii (Table 5). The mannose : glucose ratio varied from 1.41 to 5.31.

Table 5.  Solubility characteristics and mannose : glucose ratio of glucomannans (GM) in storage tissues of different geophyte species
Species/cultivar% GM soluble in water% GM insoluble in waterMannose : glucose ratio
  1. Values are means, n = 3.

Arum italicum99.50.52.24
Camassia leichtlinii ‘Caerulea’
Hyacinthus orientalis ‘Carnegie’
Hyacinthus orientalis ‘Pink Pearl’
Narcissus ‘Carlton’
Narcissus ‘Ice Follies’
Narcissus ‘Minnow’
Narcissus ‘Tete a Tete’
Scilla siberica99.40.51.87

HPAE-PAD profiles of soluble sugars and fructan

The HPAE-PAD of ethanol-soluble carbohydrates revealed unique elution profiles for each species (Fig. 2). Note that none of these chromatograms contain an internal standard. Characteristically, in the PA-1 column, mono- and disaccharides elute first, followed by longer-chain polymers (potentially glucan, fructan and any other oligosaccharides). Typically, fructan polymers began eluting after c. 10 min (under our conditions), and could be resolved to 20 or more individual peaks, depending on the species and its inherent carbohydrate profile (Fig. 2). Fructans from several geophytes resolved particularly well (e.g. Narcissus sp. up to 40 peaks and Iris bucharica up to 50 peaks). While different species exhibited different profiles, different cultivars of the same species (e.g. Tulipa gesneriana ‘Monte Carlo’ vs ‘Apeldoorn’, and H. orientalis ‘Carnegie’ vs ‘Pink Pearl’) exhibited identical profiles, although peak heights were different between cultivars. The peaks and retention times indicate the resolution of isomers of polymers with the same DP in many species. Changes in eluent strength (especially sodium acetate) remarkably changed retention times, but the general elution pattern remained the same (data not shown). Reducing sodium acetate concentration below 0.5 m resulted in incomplete elution of the longest fructan polymers (data not shown). On the other hand, shallow sodium acetate gradients (increasing 0 to 0.5 m sodium acetate over a longer duration) improved the resolution of individual peaks.

Figure 2.

Figure 2.

High-performance anion-exchange chromatography with pulsed amperometric detection (HPAE-PAD) chromatograms of ethanol-soluble carbohydrates from different geophytes. Ethanol extracts were subjected to chromatography under the conditions described in the Materials and Methods section. The elution positions of glucose, fructose and sucrose are indicated in the first chromatogram.

Figure 2.

Figure 2.

High-performance anion-exchange chromatography with pulsed amperometric detection (HPAE-PAD) chromatograms of ethanol-soluble carbohydrates from different geophytes. Ethanol extracts were subjected to chromatography under the conditions described in the Materials and Methods section. The elution positions of glucose, fructose and sucrose are indicated in the first chromatogram.

Starch solubility in water

The solubility of starch during the extraction varied depending on the species, medium of extraction and extraction temperature (Table 6). When tissues were extracted directly with water at 25°C, from 1 to 17% of total starch became soluble depending on the species. Iris species, in particular, had a high percentage of soluble starch. This soluble starch fraction could be a distinct pool of soluble alpha glucans. Boiling for a short duration is a usual practice to inactivate enzymes when carbohydrates are extracted with water. When tissues were boiled for 5 min initially before continued extraction at 25 or 70°C, a substantial portion of starch became soluble regardless of the subsequent extraction temperature. The starch solubility at 70°C was slightly higher than that at 25°C (after initial boiling) in most of the species, with the greatest effect in Camassia forbesii and Iris reticulata. The results indicate that the starch granules of most of the species may not be heat-stable when extracted in water. Extraction with water at 70°C after extraction with ethanol, however, significantly reduced starch solubility.

Table 6.  Solubility of starch from storage tissues of different geophyte species
Species/cultivar% of starch soluble in water at the indicated temperature
25°C (5 min boiling)70°C (5 min boiling)25°C (no boiling)70°C (after ethanol extraction)
  1. The percentage of starch solubilized (i.e. not retained in the residue) with different extraction methods based on the total starch determined in residue after ethanol extraction is given. Values are means, n = 3.

Arum italicum5264 811
Camassia forbesii36731214
Crocus vernus ‘Pickwick’3334 6 8
Colchicum ‘The Giant’79861519
Frittilaria imperialis ‘Rubra Maxima’2331 3 5
Fritillaria persica2831 5 7
Galanthus nivalis70861114
Hyacinthus orientalis ‘Carnegie’6567 810
Hyacinthus orientalis ‘Pink Pearl’5869 612
Iris bucharica365412 7
Iris reticulata ‘J.S. Dyt’47831512
Iris reticulata ‘Pauline’56871710
Iris xiphium ‘Blue Diamond’7282 3 9
Muscari armeniacum527211 8
Narcissus ‘Carlton’5251 3 9
Narcissus ‘Ice Follies’3433 310
Narcissus ‘Minnow’6264 110
Narcissus ‘Tete a Tete’6476 3 6
Scilla siberica5754 8 8
Tulipa gesneriana ‘Apeldoorn’8497 510
Tulipa gesneriana ‘Monte Carlo’9086 212
Tulipa tarda9197 210
Tulipa turkestanica8686 3 9

Ethanol extracts did not contain any glucose polymers as confirmed by absence of glucose release when digested with amyloglucosidase. The presence of glucose polymers in the water extracts was confirmed by the release of glucose molecules after amyloglucosidase digestion. The HPAE-PAD chromatograms of water extract did not show any additional peaks compared with the ethanol extracts, suggesting that the water-soluble glucose polymers were too large to be resolved with HPAE-PAD. Appearance of maltose or malto-oligosaccharides also could not be detected. When the water extracts (or water extracts after ethanol extraction) were frozen, and then thawed, the glucose polymers formed a white precipitate that could be completely recovered as a pellet when centrifuged in a microcentrifuge at 16 500 g for 10 min. No fructan or glucomannan was precipitated under these conditions, except in G. nivalis where > 50% of the fructan precipitated. After centrifugation, the supernatant was completely free of glucose polymers.

Based on these results, during routine total fructan and glucomannan determinations, water-extracted samples were frozen, thawed, centrifuged, and the supernatant was used for acid hydrolysis. The supernatants were also digested with amyloglucosidase to confirm the absence of soluble starch.


Nonstructural carbohydrates accounted for 50–80% of the dry weight in the geophyte storage organs investigated in this study. Starch, fructan, soluble sugars, and glucomannan were the major nonstructural carbohydrates detected in these organs. Various combinations of different nonstructural carbohydrates occurred, depending on the species, and a wide range of concentrations of each carbohydrate was evident. Most of the nonstructural carbohydrate (> 90%) was stored as polymers (starch or fructan). Starch was the most abundant nonstructural carbohydrate on a concentration basis and was absent only in Allium sp. and C. leichtlinii. The absence of starch in storage organs of Allium has been reported previously, although starch has been detected in Allium cepa stem, root tip initials and in the primary thickening meristem (Ernst & Bufler, 1994).

Fructan occurred as the exclusive polysaccharide only in Allium. In many other geophytes, fructan occurred concurrently with starch, but at a lower concentration (except in Muscari armeniacum). Also a wide range of concentrations of fructan was evident between species. The exact physiological role of fructans in plants is not yet fully understood (Vijn & Smeekens, 1999). The occurrence of fructans concurrent with starch suggests that they are not merely an alternative form of reserve carbohydrate. In a study of nonstructural carbohydrates in leaves of 185 accessions of warm- and cool-season grasses, Chatterton et al. (1989) showed that fructans accumulated mainly in cool-season grasses during cool temperatures. Fructans accumulated concurrently with starch, and under the conditions that were conducive to fructan accumulation, starch and soluble sugars also accumulated, indicating that fructans were not synthesized at the expense other forms of carbohydrates. In a study of carbohydrates in 20 native British species of angiosperms over four seasons, Broklebank & Hendry (1989) observed that fructan was associated with early-season, low-temperature growth. Orthen (2001a) conducted a survey of reserve polysaccharides in storage organs of 63 geophytic species native to the winter-rainfall region of South Africa. She reported that 24 species stored starch exclusively, 24 species stored fructan exclusively, and 15 species stored starch and fructan concurrently. Apart from the observation that all the species (except one) that stored both starch and fructan belonged to Monocotyledonae, no phenological characteristic could be attributed to the fructan accumulation. Assuming that the storage organs of all the geophytes in our study were at a comparable physiological stage, any obvious correlation cannot be made between fructan concentration and taxonomical or phenological characteristic. Many geophytes require an exposure to a period of low temperature for normal growth and flowering. Changes in fructan are among the biochemical features observed during this low-temperature phase. For example, an increase in fructan concentration during the low-temperature ‘dormancy release’ has been observed in tulips (Kamenetsky et al., 2003). Cryoprotection has been suggested as a possible role of fructans (Pontis, 1989), and fructan may be fulfilling this role in underground organs of geophytes.

Glucomannan occurs in species belonging to Liliaceae, Amaryllidaceae, Orchidacae and Araceae (Bewley & Reid, 1985; Wozniewski et al., 1991). All the glucomannans isolated from bulbs and tubers are α-(1-4)-linked, slightly branched, and water-soluble, characteristics that are different from seed mannans (Bewley & Reid, 1985). In our study, glucomannan was a far less abundant carbohydrate polymer compared with starch and fructan. The occurrence of glucomannan, reported earlier in Narcissus sp. (Tomoda et al., 1980), Arum (Achtardj & Koleva, 1973), and S. siberica (Barbakadze et al., 1996), was confirmed in our study. Only in C. leichtlinii did glucomannan constitute a significant concentration (146 mg g−1 DW). The reserve carbohydrate composition of C. leichtlinii is interesting in that it contained only fructans and glucomannan as polysaccharides with no detectable starch. In most glucomannans, mannose contributes more to the polymer than glucose. Mannose : glucose ratios reported for geophytes are 2.5 in Lilium bulbs (Tomoda et al., 1976) and 5 in Narcissus bulbs (Tomoda et al., 1980). The mannose : glucose ratios in our study ranged from 1.41 to 5.31. The ratios for Narcissus sp. in our study are similar to those reported earlier (Tomoda et al., 1980). In bulbs and tubers, glucomannans are stored in specialized cells called idioblasts (Meier & Reid, 1982). The hydrophilic nature of glucomannan suggests that it may have a function in water storage (Bewley & Reid, 1985); however, its specific functions in geophytes included in this study are not clear.

HPAE-PAD is especially useful for analysis of carbohydrate polymers (Rohrer, 2003) and can achieve successful separation of glucose polymers up to 80 DP (Hanashiro et al., 1996), and fructans up to 40 DP (Chatterton et al., 1993). The HPAE-PAD is capable of resolving polymers not only up to individual degree of polymerization but also structural isomers of oligosaccharides in the low-DP range (Bancal et al., 1993). A major drawback of HPAE-PAD, however, is that the degree of detector response is different for each carbohydrate (usually a decreasing response with increasing DP) and thus quantification is difficult, owing to a lack of accessible standards to calibrate each peak (Legnani & Miller, 2001). It has been shown that HPAE-PAD chromatograms can be used to ‘fingerprint’ fructans from different species as a result of distinct elusion profiles obtained from different species (Chatterton et al., 1990, Ernst et al., 1998). In the current study, we observed reproducible and unique chromatographic profiles for different species, whereas differentiating cultivars within a species was not possible (Fig. 2). Care should be taken, however, in using fructan elution profiles to identify plant sources, as the fructan profiles can be altered by the physiological state of the tissue. Also chromatographic conditions that affect the elution profile, such as the type and length of separating column, oxidation potentials, and strength and gradient pattern of eluents, should be clearly defined when comparing fructan elution patterns.

Our study clearly demonstrates the importance of careful choice of extraction conditions for accurate determination of nonstructural carbohydrates in storage organs, especially if coexistence of starch and fructan is suspected. Special care should be taken if carbohydrates are extracted with water, and the possibility of the presence of soluble starch in water should be investigated. The best protocol to avoid starch solubility would be sequential extraction with ethanol followed by water. These ethanol and water fractions could be analyzed separately or after combination. The insoluble residue can be used for starch determination. We demonstrated that with careful optimization of conditions, acid hydrolysis can be used for accurate determination of total fructan and glucomannan. Enzymatic methods are also available for the determination of total fructans (Anderson & Sorensen, 1999; Orthen, 2001b). Commercial enzyme preparations are available for total fructan determination in extracts where mixtures of carbohydrates are present. These methods involve degradation of nonfructan carbohydrates, first to monosaccharides and then digesting fructans with fructan hydrolases. Enzymes specific for glucomannan hydrolysis are not yet commercially available.

The amount of starch remaining in the residue after water extraction water was different depending on the species, indicating that different proportions of starch had been solubilized and removed with water. The extraction temperature seems to play a major role in this characteristic. The polymers amylose (long, unbranched glucose chains) and amylopectin (shorter, branched chains of glucose) contribute to the total structure of starch. In some starch granules, a third type, phytoglycogen (a short, very highly branched chain of glucose) has also been detected (Manners, 1985). The percentage of amylose usually ranges between 15 and 25%, with exceptions in some low-amylase starches (Noda et al., 2003). The exact ratio of amylose to amylopectin is largely determined by the genetics of the species or cultivar. When starch granules are heated in water, they swell, gelatinize, and rupture, yielding an aqueous dispersion or paste (Manners, 1985). The starch gelatinization temperature varies by species and ranges from 55 to 80°C, depending on many factors, including the amylase : amylopectin ratio. In the starch granule, amylopectin makes up the continuous network in which amylose is dispersed (Mitchell, 1977). With warm water (c. 70°C), amylose of low DP can be leached from potato starch granules (Cowie & Greenwood, 1957). The degree to which the amylose and amylopectin can be made to disperse to colloidal dimension (put into ‘solution’) depends on physical manipulations such as sharp mixing (Mitchell, 1977). Our data show up to 17% of the starch (glucose polymers) was readily soluble in water at 25°C. This faction is most likely a pool of soluble alpha-glucans. The differential starch solubility we observed in different geophytic species could be the result of differences in starch grains, including different amylase : amylopectin ratios and physicochemical properties. This is an area for further study.

In conclusion, in this report we demonstrated appropriate extraction methods and analytical techniques for accurate determination of the major nonstructural carbohydrates in storage organs of several ornamental geophytes. Results showed a wide range in terms of both the type and concentrations of carbohydrates. It will be interesting to investigate the specific functions of these carbohydrates and how they are utilized during the growth and development of individual species.


We thank Anthos, the Royal Trade Association for Flowerbulbs and Nurserystock, Hillegom, the Netherlands, for providing plant materials for this work, and for partial financial support.