Materials and Methods

Horticultural plants used in this study were grown on raised wire-mesh benches in glass-glazed greenhouses located in the Plant Health Care Greenhouse Facility at the University of Illinois (Urbana–Champaign, IL). The scientific name, common name, and cultivar of each of the plants evaluated are presented in Table 1. Fifteen to 20 plants of each species were obtained from the Ball Ornamental Company (West Chicago, IL). Plants were started from rooted plugs contained in 24 round-cell Pro-Trays (Hummert International^™, Earth City, MO). Cuttings were taken from stock plants or seedlings were grown from seed (Table 1). Plants were transplanted into Elite 600 pots (7.5 L) (ITML Ornamental Products Inc., Brantford, Canada) filled with either GP2 nursery growing medium (Midwest Groundcovers, St. Charles, IL) or with Sunshine^® LC1 growing medium (Sun Gro Horticulture^© Canada Ltd., Bellevue, WA). The GP2 nursery growing medium was comprised of peatmoss, perlite, composted pine bark, and highly processed and composted cow manure. The Sunshine^® LC1 growing medium consisted of 70% to 80% Canadian sphagnum peatmoss, perlite, dolomitic limestone, gypsum, and a wetting agent. All plants received Peter’s^® 20N–4.4P–16.6K fertilizer (Scotts-Sierra Ornamental Products, Marysville, OH) applied in a constant liquid feed program at 200 ppm nitrogen. The temperature inside the greenhouses was maintained at 24 ± 2 °C (day) and 18 ± 2 °C (night). All plants were grown under natural daylight conditions with no supplemental lighting. All plants were watered, when needed, using a municipal water source, which was tap water from Champaign–Urbana, IL.

Table 1.

Descriptive list of horticultural plant species used, including scientific name, common name, cultivar, and propagation method (starting material).

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Samples were harvested from each plant species by removing the aboveground portions (flowers, leaves, branches, and stems) and placing the plant material into a #20 brown paper bag (Commercial Bag and Supply, Des Moines, IA). Five replicates (individual plants) of each plant species were harvested and used to establish baseline silicon concentrations. The garden zinnia, sweet basil, and French marigold plants were grown for 65 d and harvested on 9 Apr. 2008. The tickseed, garden phlox, and the aster plants were grown for 73 d and harvested on 1 Apr. 2008. The meadow sage, Chinese astilbe, and coral flower plants were grown for 59 d and harvested on 1 Apr. 2008. A plant alkaline fusion technique was used to quantify the silicon concentration in each of the 10 plant species. This procedure, which is described more thoroughly in Hogendorp et al. (2011), involved tissue samples from all plant species dried in a gravimetric convection oven (Precision Scientific Group, Chicago, IL) set at 64 ± 2 °C until the plant tissue reached a constant weight. The dried plant tissue was then ground into a fine homogenous powder using a cyclone sample mill (Model 3010-030 UDY Corporation, Fort Collins, CO) fitted with a 1-mm screen (Gaines et al., 1987). The dried ground plant tissue was placed in a #7 coin envelope (8.9 × 16.5 cm) (Office Depot, Inc., Delray Beach, FL) and stored at 21 ± 2 °C. Before analysis, plant tissue samples were re-dried in the gravimetric oven set at 85 °C for 2 h, mixed thoroughly, and then weighed for dry-ashing.

Fifty milligrams of dried, ground plant tissue was placed in 20-mL nickel crucibles fitted with corresponding nickel lids (Fisher Scientific International, Fairlawn, NJ) and ashed in a muffle furnace (Hytherm Co., Pennsauken, NJ) set at 550 °C for a minimum of 4 h and then digested. Two grams of anhydrous granular sodium hydroxide was added to each nickel crucible. The covered crucible was placed on an iron wire-gauze screen and then positioned on a ring stand. A standard grid-top adjustable natural gas Bunsen burner was ignited below the nickel crucible/ring stand apparatus contained within an operating fume hood. After 5 min had elapsed, the warm nickel crucible was gently swirled using a pair of tongs, which prevented the ash from adhering to the sides of the nickel crucible. The crucible was then returned to the ring stand for 10 min (Jones and Dreher, 1996; Volk and Weintraub, 1958). The alkaline fusion process resulted in a clear liquid-molten mixture. The crucible was then cooled, resulting in a white to purple sodium–silicate fusion cake located at the bottom of the nickel crucible. Approximately 20 mL of deionized/distilled (d/d) water was added to the nickel crucible. The crucible remained idle for 6 to 8 h to dissolve the fusion cake. Finally, the silicon sample solution was transferred to a 250-mL volumetric flask and brought to volume with d/d water.

A 25-mL aliquot of the silicon sample solution was transferred to a 50-mL polypropylene beaker using a volumetric pipette. The silicon sample aliquot was acidified with 10 mL of 1N H₂SO₄ followed by the addition of 10 mL of ammonium paramolybdate tetrahydrate solution. More detailed information associated with this portion of the procedure is provided in Hogendorp et al. (2011). The silicon sample was agitated for 15 min and then remained idle for another 15 min to allow for full color development. Measurements were taken using a spectrophotometer (Model ultraviolet-160; Shimadzu Corporation, Kyoto, Japan) at 820 nm.

New colorimetric reagents were prepared every ≈7 d because the ANSA (1-amino-2-napthol-4-sulfonic acid) solution would transition from a yellow to a dull yellow–brown color and lose sensitivity if maintained too long (longer than 7 d) (Hogendorp et al., 2011). The ammonium paramolybdate tetrahydrate solution was discarded if a white precipitate formed, which commonly occurred after 10 to 12 d; however, precipitates would form sooner if the solution was acidic. A new solution of 5 N sodium hydroxide was prepared with every new ammonium paramolybdate tetrahydrate solution; otherwise, this would be a source of silicon contamination.

Results and Discussion

Zinnia and aster accumulated substantially more silicon from the municipal water source (5365 and 4796 mg·kg⁻¹ silicon, respectively) compared with the other plants evaluated, which accumulated less than 3000 mg·kg⁻¹ silicon (Table 2). These plants were grown under conditions similar to a greenhouse production system, not manipulated with purified laboratory grade water, and without a supplemental silicon-based fertilizer. As such, it is not possible to discern if the addition of supplemental silicon-based fertilizers will benefit these plant species other than increasing the overall silicon concentration. Therefore, we hypothesize that zinnia, which has an initial high silicon concentration, may benefit more from silicon-based fertilizer applications. Frantz et al. (2008) found that the silicon content of zinnia leaves was only 31 mg·kg⁻¹ when grown in purified laboratory water, but leaf tissue accumulated up to 11,750 mg·kg⁻¹ silicon (determined colorimetrically) when 2.0 mm potassium silicate fertilizer was added to the nutrient solution. This indicates that approximately half of the silicon accumulated (5,365 mg·kg⁻¹ silicon compared with 11,750 mg·kg⁻¹ silicon) in zinnia plants was fulfilled by our municipal water source and possibly the growing medium. Further studies are needed to determine if increased silicon concentrations in zinnia leaf tissue confers resistance to abiotic stresses, disease, and/or insect pests.

Table 2.

Silicon concentration (mg·kg⁻¹), sd (mg·kg⁻¹), and cv (%) of 10 commonly grown horticultural plant species using the plant alkaline fusion technique silicon quantification procedure.^z

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A plant species that may not benefit from supplemental silicon-based fertilizer applications is marigold. Frantz et al. (2008) reported that ‘African Atlantis Primrose’ marigold, Tagetes erecta L. plants, which received a nutrient solution containing 2.0 mm potassium silicate fertilizer, had 486 mg·kg⁻¹ silicon in the leaf tissues compared with 129 mg·kg⁻¹ silicon (determined colorimetrically) when plants were grown in a nutrient solution prepared with purified, laboratory-grade water without any silicon-based fertilizer, a difference of 357 mg·kg⁻¹ silicon. The French marigold plants in our study accumulated 625 mg·kg⁻¹ silicon, which was higher than the silicon-treated ‘African Atlantis Primrose’ marigold, T. erecta, plants used by Frantz et al. (2008) possibly because of the soluble monosilicic acid levels in municipal water supplies provided ample silicon for these plant species. In addition, the municipal water sources of Champaign–Urbana, IL, may have silicon levels higher than 2.0 mm silicon. Furthermore, French marigold may be more efficient in absorbing silicon from nutrient solutions than T. erecta.

The other horticultural plants evaluated in our study including coral flower (555 mg·kg⁻¹ silicon), rosemary (1053 mg·kg⁻¹ silicon), and astilbe (1411 mg·kg⁻¹ silicon) did not accumulate substantial concentrations of silicon and as such would be categorized as “silicon rejectors” according to Ma et al. (2001). We did not test whether applications of supplemental silicon-based fertilizers will increase silicon concentrations in plant tissues or if increased silicon content would benefit these plant species such as improving water relations, increasing cold-hardiness, and enhancing disease and/or insect resistance.

One study reported a number of horticultural plants including zinnia (Zinnia elegans Jacq), impatiens (Impatiens wallerana Hook. f), New Guinea impatiens (Impatiens hawkeri Bull.), verbena (Verbena ×hybrida Voss), vinca, [Catharanthus spp. (L.) G. Don], and calibrachoa (Calibrachoa × hybrid Llave & Lex.) actually respond to silicon fertilization with increased silicon contents in the leaves (Frantz et al., 2008). The plants used in that study were grown in 18 mega-ohm purified, “silicon-free” laboratory-grade water. However, it is not apparent if municipal or well-water sources, commonly used in greenhouse production systems, would supply enough silicon to reach luxury status on their absorption curves (Marschner, 1995) or if additional supplemental silicon-based fertilizers are required.

It is important to determine if applications of silicon-based fertilizers to horticultural plants will provide benefits associated with increased heat tolerance, drought tolerance, cold-hardiness, and enhanced resistance to fungal infections and/or insect infestations. All these benefits have been affiliated with elevated silicon contents in specific agricultural crops such as rice, sugarcane, and certain cereals (Blackman, 1968; Jones and Handreck, 1967; Nayar et al., 1975). However, studies have shown that applications of silicon-based fertilizers to coleus [Solenstemon scutellarioides (L.)], fiddleleaf fig (Ficus lyrata Warb.), and poinsettia (Euphorbia pulcherrima Willd. ex Klotzsch) did not provide resistance against the citrus mealybug (Planococcus citri Risso) and greenhouse whitefly (Trialeurodes vaporariorum Westwood) (Hogendorp et al., 2009a, 2009b, 2010). In addition, Ranger et al. (2009) reported that nutrient solutions amended with potassium silicate did not affect the pre-reproductive period and survival of the green peach aphid (Myzus persicae Sulzer) reared on Z. elegans, whereas total cumulative fecundity and the intrinsic rate of increase were slightly reduced; however, this was only a modest increase in resistance levels.

Frantz et al. (2008) reported substantial differences in the silicon concentration of certain plant species when comparing the inductively coupled plasma optical emission spectrometry (ICP-OES) silicon determination procedure used by the U.S. Department of Agriculture–Agricultural Research Service laboratory that uses a separate colorimetric silicon determination procedure. However, there was no explanation to account for the differences in the silicon concentrations obtained between the silicon determination procedures. The plant species that differed substantially in silicon concentrations included snapdragon (Antirrinum majus L. ‘Rocket White’), impatiens (Impatiens wallerana Hook. f. ‘Super Elfin White’), New Guinea impatiens (Impatiens hawkeri Bull. ‘Sonic Light Lavender’), and poinsettia (E. pulcherrima ‘Freedom Red’). It was suggested that digestion of certain plant species may result in unknown compounds in the matrices, which could significantly affect the quantification procedure (Jonathan Frantz, personal communication). However, it is not known if these matrices interfere with the ICP-OES or the colorimetric procedure, and as such, further investigation is warranted. It is difficult to ascertain if the addition of supplemental silicon-based fertilizers will benefit these plant species, although it is possible that zinnia, which already has a high silicon concentration, may profit the most from silicon-based fertilizer applications.

In conclusion, the current study is just one of a few to quantify the silicon concentration in a variety of different horticultural crops. Silicon determinations are an initial step toward evaluating whether any of these plants benefit from applications of silicon-based fertilizers in regard to promoting cold-hardiness and/or plant resistance to fungal pathogens and/or insect pests. Therefore, further research needs to be conducted on the effect of silicon on the performance and resistance of horticultural crops.