DIEL PATTERNS IN VOLATILES RELEASED BY MECHANICALLY-DAMAGED WHEAT PLANTS

Dariusz Piesik¹, David K. Weaver², Gavin E. Peck², Wendell L. Morrill^2
1 Department of Applied Entomology, University of Technology and Agriculture, Bydgoszcz, Poland
² Department of Land Resources and Environmental Sciences of Montana State University, USA

ABSTRACT

Plants emit volatile chemicals, which may attract insect herbivores and associated predators or parasitoids. Insect feeding or mechanical injury may influence the release of these chemicals. Five types of plant wounding were imposed after the Zadoks 32 growth stage was reached and volatiles were collected from the main stem. The volatile collection sequence used was two consecutive ten-hour collections to obtain the samples in either daylight or darkness. The first sampling period was immediately after the injury, followed by sampling at 2 and 4 days. The volatiles collected were analyzed by coupled gas chromatography-mass spectrometry (GC-MS). Damage to wheat stems or leaves increased the production of the secondary metabolites, linalool (C₁₀H₁₈O) and linalool oxide (C₁₀H₁₈O₂). The amount released was greater during the day, and was also influenced by the type of the injury. In some cases, enhanced volatile chemical production continued for 4 days following the injury.

Key words: wheat, Triticum aestivum, volatiles, semiochemicals, odors.

INTRODUCTION

Plants have an arsenal of physical and chemical defenses against herbivores [18]. Volatile emissions from injured plants may attract herbivores [1,5,7] and their parasitoids [4,5,7]. An understanding of the factors involved in the production of volatile secondary metabolites may help to increase the effectiveness of biological control methods [13,19,23].

Volatile compounds produced by wheat, Triticum aestivum L., have been collected and identified using several techniques [3,10]. These compounds are in part responsible for the characteristic plant odor [3]. The principal volatile compounds found in wheat and oat seedlings were characterized for attractancy to insect herbivores, but wheat infested by insects also produced different volatile signals as a result of infestation [17]. Induced chemicals may influence the level of plant damage. For example, laboratory studies showed that mixed populations of aphids Sitobion avenae F. and Rhopalosiphum padi L., probed the tissue less, demonstrated longer ingestion times and an increase fecundity on wheat seedlings from pure colonies as a result of specific odors involved [12]. The present experiment was designed to measure the production of two plant-produced volatile chemicals, linalool and linalool oxide, that are known to affect insect behavior. Previous observations suggested that volatile composition and abundance might be different during day and night collections. In particular, volatile production in daylight and under darkness after several types of mechanical plant injury was compared. Based on previous studies, it is well-known that volatile production by plants is typically greater during daylight when compared to dark conditions. Moreover, different kinds of injury may cause varied reactions in both affected and unaffected plant tissues. Based on these assumptions, the following hypotheses were formulated:

• there are different amounts of linalool and linalool oxide released following different types of the injury,

• daylight may enhance the release of volatiles, while volatile production is much lower at night,

• additionally, interactions between injury types, collection dates and light conditions can be statistically verified.

Based on the previous study, it is well-known that volatile release varies over the course of the day under light conditions. Usually the largest amounts are released in the morning, and decreased in the afternoon [16]. This study was conducted to quantify the total production of these volatile compounds throughout photophase and scotophase for wheat plants that had experienced simulated defoliation or were left intact. The quantification of the amounts of volatiles produced should provide a useful range of concentrations for the development of synthetic semiochemical lures designed to mimic growing intact and injured wheat plants.

MATERIAL AND METHODS

‘McNeal’ spring wheat seeds were sown in a greenhouse with a photoperiod of 16L: 8D. The daytime temperature was 22 ± 2°C and the overnight temperature was 18 ± 2°C. Plants were grown at a density of two per pot in equal parts of sterilized silt loam soil, washed sand, Canadian sphagnum peat moss, and Sunshine Mix 1 (Canadian sphagnum peat moss, perlite, vermiculite, and Dolmitic lime – Sun Gro Horticulture, Inc., Bellevue, Washington, U.S.A.). The plants were watered four times weekly, and fertilized with Peters® General Purpose Fertilizer (J.R. Peters Inc., Allentown, Pennsylvania, U.S.A.) at 100 ppm in aqueous solution twice a week once the plants had reached the third-leaf stage.

Five types of plant wounding were selected after the Zadoks 32 growth stage was reached (stem elongation had separated the first two nodes). These were 1) pierced stem (PS) in which the main stem was punctured in five places with a 0.34 mm diameter needle, 2) scraped stem (SS) lumen in which the stem interior was penetrated with a 1.64 mm diameter needle and scraped against the opposing stem wall, 3) removal of the distal half of the upper leaf (THCL), 4) removal of the upper one-fourth of the leaf (TQCL), and 5) removal of the lower quarter of the leaf (BQCL). Scissors were used to remove leaf portions. These damage patterns were imposed to simulate various forms of insect feeding injury.

The volatile collection system (Analytical Research Systems, Inc., Gainesville, Florida, USA) consisted of six glass collection chambers, which were open at one end to enclose the growing plant. A flexible Teflon^® sleeve was tape-sealed around the base of the main stem to prevent the collection of excess soil volatiles. The chambers were 40 mm in diameter and 800mm long. Volatiles were collected simultaneously from all the six chambers. Each volatile collection chamber was fitted with manifold with 8 ports, fitted with threaded air inlet caps, and with threaded volatile collector ports, both with no. 7 ChemThread inlets (inner diameter 6.35 mm), sealed using rubber O-rings. A volatile collector trap (6.35 mm in OD, 76 mm-long glass tube; Analytical Research Systems, Inc., Gainesville, Florida, USA.) containing 30 mg of Super-Q (Alltech Associates, Inc., Deerfield, Illinois, USA) adsorbent was inserted into each port and sealed by the O-ring/ChemThread assembly. Purified, humidified air was delivered at a rate of 1.0 liter per min over the plants, and the flow and pressure were maintained by a vacuum pump. The volatile collection system was computerized and programmed with software inputs, which allowed two event controllers to switch solenoid switches off and on. These switches allowed the airflow of entrained volatiles to be switched from one port to another. This capability allowed for the programming of multiple collections from each plant during either photophase or scotophase.

Volatiles were collected from the main stem and three large uppermost leaves of each plant only. The volatile collection sequence involved two consecutive ten-hour collections to obtain the day/night samples. The first sampling period was immediately after the injury, followed by the second sampling period after 2 and again after 4 days. There were three temporary replicates of each plant and wounding type. For each collection interval, five plants were collected: three random treatment replicates and two controls. Specific treatment replicates for collection were assigned randomly each day and experiments were staged daily until the completion of the experiment. Additionally, one control chamber was collected each day. This control consisted of the airspace above the pot containing soil only.

Volatile chemicals were eluted from the Super-Q in each collection trap with 225 ľl of hexane. Then, 7 ng of decane was added as an internal standard. Volatiles were analyzed by coupled gas chromatography-mass spectrometry (GC-MS). The GC was an Agilent Technologies 6890 instrument fitted with a 30-m DB-1MS capillary column (0.25-mm-ID, 0.25 µm film thickness; J & W Scientific, Folsom, California). The temperature program increased the chromatography oven temperature from 50°C to 280°C at 10°C·min^-1. The MS instrument was an Agilent Technologies 5973. The identification of volatiles was verified with authentic standards purchased from commercial sources that had the same GC retention times and mass spectra.

The amounts of linalool and linalool oxide for each plant at each collection interval were variance analyzed. The independent variables included the injury type (Injury), days after injury (0, 2, or 4 days), and collection interval in each day/night – each consecutive ten-hour interval (Collection time). Mean amounts were separated after analysis of variance using Tukey’s test for significant differences at α = 0.05.

RESULTS

The volume of linalool and linalool oxide produced by plants was significantly higher during the day collection period than during the night. Damage types and collection time affected volatile emissions. Injured plants generally produced more chemicals than the control plants. On average, significantly damaged wheat plants released higher amounts of these terpenoids.

Overall, greater amounts of linalool (Table 1) and linalool oxide (Table 2) were released following a mechanical injury, as compared with the control. A significant amount of variation was explained by the injury type for linalool (F = 7.4, DF = 5) and linalool oxide (F = 3.5, DF = 5). Within injury types, the two quarter cut leaf treatments and ‘SS’ demonstrated significantly greater amounts of linalool than the control (Table 1), although all the injury types showed greater amounts of this compound, as compared with the control. There was found a significant interaction between the injury types and the collection time for linalool. The significant difference for the interaction was 20.5 ng for linalool, which was significantly greater for the ‘TQCL’ (40.7 ng), when compared to the four other damage types (Table 1) for the first collection period. However, on the second day, the linalool release was quite large for both the ‘BQCL’ and the ‘TQCL’ establishing the expected similarities between the two injury types. On average, these two kinds of injury caused the release of 45 ng of linalool, a 10-fold greater amount than for the other injury types. Generally the greatest volatile production was observed on the first day of collection time, except for the two quarter cut leaf treatments. In contrast, the ‘SS’ and both quarter cut leaf injury types caused the release of significantly greater amounts of linalool oxide, while all injury types released greater amounts only (Table 2).

Table 1. Amount of linalool collected from mechanically-injured and control wheat plants, ng

Collection time (II)		Injury (I)
		Control	PS	SS	THCL	TQCL	BQCL	Mean
0 day	0-10 h day	6.38	13.04	24.39	10.18	40.66	23.73	19.73
	0-10 h night	0.21	0.49	5.38	0.90	1.08	0.00	1.35
2nd day	0-10 h day	1.90	1.93	7.74	6.12	52.39	41.47	18.59
	0-10 h night	0.00	0.00	1.71	0.38	0.00	0.00	0.35
4th day	0-10 h day	1.85	1.62	18.56	3.38	7.87	4.17	6.24
	0-10 h night	0.00	0.00	0.93	0.00	0.00	1.38	0.38
Mean		1.72	2.85	9.79	3.49	17.00	11.79	7.77

LSD 0.05 for injury (I) – 8.34 LSD0.05 for collection time (II) – 8.34 LSD0.05 for I/II – 20.49 LSD0.05 for II/I – 20.49

Table 2. Amount of linalool oxide collected from mechanically-injured and control wheat plants, ng

Collection time (II)		Injury (I)
		Control	PS	SS	THCL	TQCL	BQCL	Mean
0 day	0-10 h day	0.74	1.48	5.73	2.02	5.99	5.26	3.54
	0-10 h night	0.38	0.00	4.51	1.10	1.51	0.00	1.25
2nd day	0-10 h day	0.00	0.00	1.18	0.00	23.29	12.48	6.16
	0-10 h night	0.00	0.00	1.71	1.09	1.61	0.00	0.74
4th day	0-10 h day	0.00	0.00	5.00	1.98	4.48	3.74	2.53
	0-10 h night	0.00	0.00	0.00	0.00	0.26	0.50	0.13
Mean		0.19	0.25	3.02	1.03	6.19	3.66	2.39

LSD0.05 for injury (I) – 4.68 LSD0.05 for collection time (II) – 4.68 LSD0.05 for I/II – non-significant LSD0.05 for II/I – non-significant

No significant interaction for linalool oxide was evident between the collection time and the injury type. The variance contributions for both linalool and linalool oxide showed significant interactions across some explanatory variables, indicating that the effects of the collection day and the collection interval for each day varied by injury type (Table 3).

Table 3. The contribution of each of the classification variables to the variation in the amount of linalool and linalool oxide produced by mechanically-injured wheat plants

Linalool	SS	DF	MS	F	p
Intercept	6527.424	1	6527.424	72.58159	0.000000
Injury	3321.160	5	664.232	7.38592	0.000012
Collection time	7441.527	5	1488.305	16.54919	0.000000
Interaction	7230.809	25	289.232	3.21612	0.000058
Error	6475.120	72	89.932

Linalool oxide	SS	DF	MS	F	p
Intercept	616.811	1	616.8112	21.83693	0.000013
Injury	499.312	5	99.8624	3.53542	0.006487
Collection time	444.599	5	88.9197	3.14802	0.012599
Interaction	1129.770	25	45.1908	1.59989	0.063431
Error	2033.729	72	28.2462

A significant portion of the variation in the amount of linalool (F = 16.5, DF = 5) and linalool oxide (F = 3.1, DF = 5) was explained by the collection time. The collections for each photophase obtained significantly more volatiles than the night collection intervals. On average, linalool was released by mechanically injured wheat plants in photophase at levels that ranged from 14- to 53-fold higher than collections at night. A similar, lower magnitude of increase was also apparent for the release of linalool oxide.

Some other patterns in volatile release were apparent from these experiments. Linalool and linalool oxide release by ‘TQCL’ plants were clearly highest across the collection intervals on the third day after the injury, peaking at a mean of 52.4 ng, 23.3 ng, respectively. For both the ‘PS’ and ‘THCL’ damage types, no significant change in linalool release was recorded for any collection time. A similar pattern was evident for the ‘THCL’ treatment release of linalool oxide.

The third day in the collection times, four days post-injury, yielded less volatiles. Furthermore, at night the injured wheat plants released detectable amounts of volatiles, but in very small quantities, when compared to the release during photophase. On average, a 3-fold higher amount of linalool was released on the 1^st and 2^nd photophase of the collection times to compare with the 3^rd one. This pattern was also observed for linalool oxide.

DISCUSSION

Plants can produce secondary metabolites, which act as signals modifying the development or behavior of other organisms without showing a direct physiological activity [6], although these compounds may also play a role in defense against pathogens. Also plant tissues may show chemical changes following damage [2]. Hoballah and Turlings [11] reported on any type of surface damage commonly making plant leaves release green leaf volatiles. In the present experiment the abundance of linalool and linalool oxide collected was higher for injured plants. Generally, it was noted that different types of mechanical injuries caused the release of greater amount of potential secondary metabolites in elongating wheat plants. The increases in linalool and linalool oxide were quite distinct depending on the injury type. They were also quite distinct depending on the time after the injury. Systemically released compounds like linalool are known to be induced by caterpillar feeding damage, slightly increased by a mechanical injury, and are not released in significant amounts by undamaged cotton plants [22]. Koschier et al. [14] demonstrated that western flower thrips (Frankliniella occidentalis) were attracted by linalool. Rodriguez-Saona et al. [21] revealed that cotton plants treated with exogenous methyl jasmonate emitted elevated levels of linalool to those similar to plants damaged by herbivores, showing that this compound is an inducer of this secondary metabolite. The effects observed encourage further investigation of the ecological role of these active substances [15]. It might be expected that mechanically injured wheat plants would produce volatiles in higher amounts in the daylight as compared with the night amounts. However, the differential patterns in maximal production of these compounds across injury types and over time merit further investigation.

It is of importance to investigate how these secondary metabolites can impact natural enemy and herbivore behavior [24]. Engelberth et al. [8] demonstrated a specific function of green leafy volatiles (GLV) against herbivorous insects. Moreover, tomato plants (Lycopersicon esculentum) in response to insect feeding, release both locally and systemically elevated levels of volatile organic compounds [9]. Reddy and Guerrero [19] suggested that host plants could affect the insect physiology and behavior, including reproduction. However, volatiles emitted from plants in response to insect damage can vary with insect feeding habitats [20].

CONCLUSIONS

1. Mechanical injury of wheat plants resulted in the collection of greater amounts of the secondary metabolites: linalool and linalool oxide. Moreover, the greatest collection of volatiles occurred during daytime or in photophase.

2. Night collection of volatiles was characterized by a lower production of these potential semiochemicals, and the small amounts of the material collected might be possibly the final carryover of photophase-driven volatile release.

3. Surprisingly, statistical analysis of the interactions showed that similar injury types caused different temporary patterns in the increased production of these volatiles.

ACKNOWLEDGMENTS

The USDA/CSREES Special Research Grant entitled “Novel Semiochemical– and Pathogen–Based Management Strategies for Wheat Stem Sawfly and Cereal Aphids” supported this project. The Montana Agricultural Experiment Station provided additional support. Dariusz Piesik is grateful to the Batory Organization for help while stay in the USA.

REFERENCES

1. Agrawal A.A., Tuzun S., Bent E., 1999. Induced plant defenses against pathogens and herbivores. APS Press, St Paul, Minnesota.

2. Banchio E., Zygadlo J., Valladares G.R., 2005. Effects of mechanical wounding on essential oil composition and emission of volatiles from Minthostachys mollis. J. Chem. Ecol. 31(4), 719-727.

3. Buttery R.G., Xu Cheng-Ji, Ling L.C., 1985. Volatile components of wheat leaves (and stems): possible insect attractants. J. Agric. Food Chem. 33, 115-117.

4. Cardoza Y.J., Albron H.T., Tumlinson J.H., 2002. In vivo volatile emissions from peanut plants induced by simultaneous fungal infection and insect damage. J. Chem. Ecol. 28, 161-174.

5. Cardoza Y.J., Teal PEA, Tumlinson J.H., 2003. Effect of peanut plant fungal infection on oviposition preference by Spodoptera exigua and on host-searching behavior by Cotesia marginiventris. Environ. Entomol. 32(5), 970-976.

6. Chamberlain K., Pickett J.A., Woodcock C.M., 2000. Plant signaling and induced defence in insect attack. Mol. Plant Pathol. 1, 67-72.

7. De Moraes C.M., Mescher M.C., Tumlinson J.H., 2001. Caterpillar-induced nocturnal plant volatiles repel nonspecific females. Nature 410, 577-580.

8. Engelberth J., Alborn H.T., Schmelz E.A., Tumlinson J.H., 2004. Airborne signals prime plants against insect herbivore attack. Plant Biology 101(6), 1781-1785.

9. Farag M.A., Paré P.W., 2002. C₆ – green leaf volatiles trigger local and systemic VOC emission in tomato. Phytochemistry 61, 545-554.

10. Hatanaka A., 1993. The biogeneration of green odour by green leaves. Phytochemistry 34, 1201-1281.

11. Hoballah M.E., Turlings T.C.J., 2005. The role of fresh versus old leaf damage in the attraction of parasitic wasps to herbivore-induced maize volatiles. J. Chem. Ecol. 31(9), 2003-2018.

12. Johansson C., Pettersson J., Niemeyer H.M., 1997. Interspecific recognition through odours by aphids (Sternorrhyncha: Aphididaea) feeding on wheat plants. Eur. J. Entomol. 94, 557-559.

13. Kessler A., Baldwin I.T., 2001. Defensive function of herbivore-induced plant volatile emissions in nature. Science 291, 2141-2144.

14. Koschier E.H., De Kogel W.J., Visser J.H., 2000. Assessing the attractiveness of volatile plant compounds to western flower thrips Frankliniella occidentalis. J. Chem. Ecol. 26, 2643-2655.

15. Ninkovic V., Ahmed E., Glinwood R., Pettersson J., 2003. Effects of two types of semiochemical on population development of the bird cherry oat aphid Rhopalosiphum padi in a barley crop. Agr. Forest Entomol. 5, 27-33.

16. Piesik D., Weaver D.K., Peck G.E., Morrill W.L., 2006. Mechanically-injured wheat plants release greater amounts of the secondary metabolites linalool and linalool oxide. J. Plant Prot. Res. 46(1), 29-39.

17. Quiroz A., Niemeyer H.M., 1998. Activity of enantiomers of sulcantol on apterae of Rhopalosiphum padi. J. Chem. Ecol. 24, 361-370.

18. Rasmann S., Köllner T.G., Degenhardt J., Hiltpold I., Toepfer S., Kuhlmann U., Gershenzon J., Turlings T.C.J., 2005. Recruitment of entomopathogenic nematodes by insec t-damaged maize roots. Nature 434, 732-737.

19. Reddy G.V.P., Guerrero A., 2004. Interactions of insect pheromones and plant semiochemicals. Trends Plant Sci. 9, 253-261.

20. Rodriguez-Saona C., Crafts-Brandner S.J., Caňas L.A., 2003. Volatile emissions triggered by multiple herbivore damage: beet armyworm and whitefly feeding on cotton plants. J. Chem. Ecol. 29(11), 2539-2550.

21. Rodriguez-Saona C., Crafts-Brandner S.J., Paré P.W., Henneberry T.J., 2001. Exogenous methyl jasmonate induces volatile emission in cotton plants. J. Chem. Ecol. 27, 679-695.

22. Röse U.S.R., Lewis J.W., Tumlinson J.H., 1998. Specificity of systemically released cotton volatiles as attractants for specialist and generalist parasitic wasps. J. Chem. Ecol. 24, 303-319.

23. Thaler J.S., 1999. Jasmonate-inducible plant defenses cause increased parasitism of herbivores. Nature 399, 686-688.

24. Wang Q,H., Dorn S., 2003. Selection on olfactory response to semiochemicals from a plant-host complex in a parasitic wasp. Heredity 91, 430-435.

 

Accepted for print: 12.10.2006

---

Dariusz Piesik
Department of Applied Entomology,
University of Technology and Agriculture, Bydgoszcz, Poland
20 Kordeckiego St., 85-225 Bydgoszcz, Poland
email: dpiesik@interia.pl

David K. Weaver
Department of Land Resources and Environmental Sciences of Montana State University, USA
334 Leon Johnson Hall, Bozeman, MT 59717-3120, USA

Gavin E. Peck
Department of Land Resources and Environmental Sciences of Montana State University, USA
334 Leon Johnson Hall, Bozeman, MT 59717-3120, USA

Wendell L. Morrill
Department of Land Resources and Environmental Sciences of Montana State University, USA
334 Leon Johnson Hall, Bozeman, MT 59717-3120, USA

---

Responses to this article, comments are invited and should be submitted within three months of the publication of the article. If accepted for publication, they will be published in the chapter headed 'Discussions' and hyperlinked to the article.

---

Main  -  Issues  -  How to Submit  -  From the Publisher  -  Search  -  Subscription