Physiology and growth of Douglas-fir seedlings treated with
ethanol solutions
Gladwin Joseph
a, Rick G. Kelsey
b,*
aDepartment of Forest Science,Oregon State Uni6ersity,Cor6allis,OR97331,USA
bUSDA Forest Ser6ice,Pacific Northwest Research Station,3200Jefferson Way,Cor6allis,OR97331,USA
Received 17 May 1999; received in revised form 14 September 1999; accepted 14 September 1999
Abstract
Applying 1, 5, 10, and 20% solutions of ethanol to the roots of Douglas-fir (Pseudotsuga menziesii [Mirb.] Franco) seedlings three times a week was deleterious to their physiology and growth. Ethanol concentrations of 10% or higher were lethal within a week of treatment initiation, while the 5% solution was lethal to seedlings at 8 weeks. Seedlings treated with the 1% solution were alive at 8 weeks, but showed signs of physiological decline. If Douglas-fir seedlings have a tolerance threshold for ethanol solutions applied to their roots, it appears to be at a concentration below 1%. Ethanol moved up the stems and into needles, yielding concentrations in the stems 9 times higher than in needles. Ethanol vapors in the atmosphere surrounding seedlings readily diffused into needles, but not into stems. After 1 week of treatments, net photosynthesis, stomatal conductance, and transpiration declined as ethanol concentrations increased. However, seedlings treated with the control (0%) and 1% ethanol solutions had the same xylem water potentials, which were higher than for seedlings treated with the 5% solutions. High ethanol concentrations (]1%) may have damaged membranes involved in photosynthesis and stomatal function thereby causing the observed decline in net photosynthesis and stomatal conductance. At concentrations]5%, water uptake was impaired, suggesting that root membranes may have been damaged. © 2000 Elsevier Science Ireland Ltd. All rights reserved.
Keywords:Pseudotsuga menziesii; Gas exchange; Tolerance; Water potential; Toxicity
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1. Introduction
Ethanol is a product of anaerobic metabolism that occurs naturally in plant tissue in response to hypoxia or anoxia [1]. Ethanol synthesis may al-low tissues to survive transient periods of oxygen stress by supplying ATP [2], and by preventing cellular acidosis [3,4]. Ethanol is produced in roots of flooded plants and diffuses into the surrounding water and soil [5], or is transported into the stem and foliar tissues where it may be metabolized into cellular constituents [6 – 8]. However, ethanol can be toxic if it accumulates to high enough concen-trations either in the rooting solution or in the tissues [9,10].
The effects of ethanol on plant growth and physiology are not consistent. It can either damage or stimulate growth depending on the concentra-tions, species, and tissue. When tomato roots were fed 5% ethanol solution growth was reduced, and solutions ]10% were lethal [11]. In contrast, fo-liar applications of 15 – 20% ethanol solutions in-creased growth of tomato (Lycopersicum esculentum Mill.) by 18% [11]. Foliar applications of 1 – 10% ethanol solutions neither stimulated growth nor caused any injury to Douglas-fir and ponderosa pine (Pinus ponderosa (Dougl.) Ex Laws.) seedlings [12]. Mortality of chick pea (Cicer arietinum L.) seedlings increased as the length of exposure to ethanol vapors increased in a static anaerobic atmosphere [13]. Cell growth and so-matic embryogenesis of carrot (Daucus carota L.) cell cultures were negatively affected at relatively
* Corresponding author. Tel.:+1-541-750-7368; fax:+ 1-541-750-7329.
E-mail address:[email protected] (R.G. Kelsey)
low concentrations (0.05%) of ethanol [10]. How-ever, placing etiolated cuttings from mung bean (Vigna radiataL.) into 0.1% ethanol solution stim-ulated root initiation and growth [14]. Root growth also was enhanced by 50% when excised wheat (Triticum aesti6um L.) roots were treated
with 0.9% ethanol in light [15]. Pea plants were able to tolerate ethanol applied to their roots at 100 times the concentration (0.46%) found in their xylem sap during flooding [9]. When cut stems of Douglas-fir seedlings were supplied with ethanol concentrations (0.1%) 2 – 3 times the amount in their stems under flooded conditions, there was no effect on stomatal conductance [5].
In our continuing effort to better understand the physiological and ecological implications of ethanol synthesis and accumulation in conifers we were interested in determining how ethanol ap-plied to the roots would affect growth and physi-ology of Douglas-fir seedlings. Since foliage of Douglas-fir is apparently capable of metabolizing ethanol [5], we hypothesized that shoot biomass might increase with some concentrations of ethanol. Additionally, by using a wide range of ethanol concentrations we attempted to establish a tolerance threshold for ethanol applied to Dou-glas-fir roots. Finally, we hypothesized that detri-mental concentrations of ethanol would affect membrane bound processes such as photosynthesis [18], stomatal conductance (via guard cell metabolism), and water uptake because ethanol toxicity is potentially associated with membrane damage [9,16,17]. Since guard cell metabolism is linked to membrane bound processes [19], stom-atal conductance would decline if ethanol damages the guard cell membrane. A decline in water po-tential in ethanol treated seedlings compared to controls under similar vapor pressure deficits would indicate damage to roots and a reduction in water uptake.
2. Material and methods
Douglas-fir seedlings were germinated and grown at Champion International Co. Nursery, Lebanon, OR. Seeds (zone 422, Central Washing-ton, elev. 610 m) were sown on 23 April, 1993, in styroblocks (192 cavities with 131 cm3 volume
each) containing a peat:vermiculite mixture (3:2). They were irrigated and fertilized twice each week
until moved to a greenhouse on 26 July. Under these conditions the seedlings had started to flush prior to initiating treatments on 27 July. These seedlings measured 4.3 cm from the root collar to stem apex, with shoot biomass averaging 0.16 g dry weight. The greenhouse was maintained at 25:15°C day and night temperature. The photo-synthetic photon flux density (PPFD) during the day was 300 – 450mmol m−2s−1. The photoperiod was extended to 16 h with sodium vapor lamps (PAR 145 mmol m−2 s−1 at plant height).
In the greenhouse each styroblock was broken into four smaller styroblocks with each containing 48 seedlings. Fifteen such styroblocks were ran-domly assigned among three experimental blocks, each with one treatment of 0, 1, 5, 10, or 20% aqueous ethanol (100%, v/v) in a 0.2% nutrient solution (15:35:15 N:P:K, Miracle-gro®). The five
styroblocks in each block were arranged in a circle and elevated off the bench to prevent cross-con-tamination from drainage.
Seedlings were watered with their respective nu-trient-ethanol solutions three times per week. A total of 4 l of treatment solution were poured into a plastic container (40.6×28×23 cm, 21.5 l) and the appropriate styroblock submerged about halfway in the solution. After 20 min they were drained and returned to their bench positions. From a preliminary experiment we determined that 20 min was sufficient for seedlings to absorb ethanol through the roots. Seedlings were treated with ethanol from 27 July to 21 September (8 weeks), then the experiment was terminated.
(Hewlett-Packard 5890, Palo Alto, CA) with a J&W Scientific DB-WAX column, 30 m×0.32 mm i.d. and a 0.25-mm film thickness. Instrument settings were as described previously [5]. Samples were analyzed by multiple headspace extraction with two injections per vial and venting between injections [20]. The instrument was calibrated with vials containing a 5-ml ethanol standard diluted with water. After analysis, tissue samples were oven dried at 102°C for 16 h, cooled in a desicca-tor for 30 min, and weighed. Ethanol concentra-tions in the tissue were calculated from headspace concentrations [20].
One day (24 h) after the third ethanol treatment (i.e. 1 week after initiating the experiment), gas exchange (net photosynthesis, stomatal conduc-tance, transpiration), xylem pressure potential (XPP), and ethanol concentrations were measured. Gas exchange measurements were made on 2 seedlings per block for the 0, 1 and 5% ethanol treatments. By this time, seedlings from the 10 and 20% ethanol treatments were showing visible signs of severe foliar and stem injury and therefore were not used. Gas exchange measurements were taken at 06:40, 10:40, 14:40, 18:30, and 22:30 h (3 h after the lights were turned off) with a LI-COR 6250 portable infrared gas analyzer (LI-COR Inc., Lin-coln, NE). The gas exchange measurement at
22:30 was made in the dark and provided an estimate of dark respiration. Gas exchange mea-surements were made on older needles after re-moving the newer expanding needles. Wound respiration was considered negligible because the cut surface area of new needles was a small frac-tion of the total surface area of old needles. After the measurements, needles were harvested and their areas estimated to the nearest 0.01 cm2 with
a video camera connected to a computer with image analyzing software (Agvision, Decagon Devices Inc., Pullman, WA). Xylem pressure po-tential was measured with a pressure chamber (PMS Co., Corvallis, OR) at predawn and again at midday on two randomly selected seedlings from each block of 0, 1, and 5% treatments. Ethanol also was quantified as before on old needles and stems of two seedlings from each block of 0, 1, and 5% treatments at 07:30 and 15:30 h.
One day (24 h) after the last ethanol application (8 weeks after treatment initiation), XPP, gas ex-change, and ethanol concentrations were measured as described above at 09:30, 10:45, and 12:00 noon, respectively, on two seedlings per block for the 0, 1, and 5% ethanol treatments. Shoot length and dry weight also were measured on 4 – 10 seedlings per block. Shoot lengths were measured from the root collar to stem apex. Shoot biomass above the root collar was determined gravimetri-cally after drying at 70°C for 48 h. These final measurements were restricted to seedlings that were still alive and physiologically functional to ensure that we measured sub-lethal effects of ethanol. This was only a concern in the 5% treat-ment where many of the seedlings had died or were dying. Seedling mortality was not recorded periodically because it was difficult to determine if seedlings were dead using visual symptoms.
Needles of control seedlings positioned next to ethanol-treated seedlings in the main experiment above showed small increases in ethanol during the first 2 h after treatments were imposed (Fig. 1), indicating ethanol vapors from the surrounding atmosphere had diffused into the control needles. To examine this more closely, on 30 August three untreated styroblocks (0%) were placed on a bench 4 – 5 m away from seedlings in the main experi-ment to ensure the atmosphere was relatively free of ethanol vapors. These untreated blocks were all watered with 0% ethanol for 20 min and then two
seedlings randomly harvested from each block. Their needles and stems were sampled as above for ethanol analysis at 0, 1, 2, and 4 h after treatment initiation. These samples served as the unexposed-controls. On the following day, three styroblocks used for the 20% ethanol treatment in the main experiment above (now with dead seedlings) were placed alternately between the 0% controls and each watered with their respective ethanol solu-tions for 20 min. Two seedlings from the 0% treatment were harvested and their needles and stems sampled for ethanol analysis at 0, 1, 2, and 4 h after treatment initiation, as described previ-ously. This was the exposed-treatment. The atmo-sphere surrounding these seedlings also was sampled for ethanol at each time. One air sample was taken at seedling height from both ends of the three styroblocks. Air (30 cm3) was drawn with a
plastic syringe and injected into a sealed autosam-pler vial with a needle inserted through the septum serving as an outlet for the displaced air from the vial. The outlet and inlet needle from the syringe were immediately removed from the septum. Ethanol in the sealed vial was analyzed as before. All statistical analyses were made with SAS software [21] using styroblocks as the experimental unit, with a single mean value for each parameter obtained from subsamples within blocks. Ethanol concentrations measured 24 h after the first treat-ment, and after 1 week (three treatments) were analyzed separately for needles and stems as a split-plot design with treatment as the main plot and time as the sub-plot. Similarly, the experiment on ethanol absorption by needles was analyzed as a split-plot design with the unexposed-control and exposed-treatment as main effects and time as the sub-plot. Diurnal gas exchange after 1 week was analyzed as a split-plot design with treatment as the main plot and time as a repeated measure, because these measurements were repeatedly made on the same seedlings. Xylem pressure potentials measured at 24 h, 1 week, and 8 weeks were analyzed separately for each time as one-way ANOVAs with treatment as the main effect. Dark respiration (after 1 week), and the final measure-ments (after 8 weeks) for shoot length, dry weights, gas exchange, and tissue ethanol concen-trations were each analyzed as a one-way ANOVA with treatment as the main effect. Where neces-sary, data were natural log transformed to meet homogeneity of variance, and normality
assump-tions. Geometric (back transformed) means are presented for data that was transformed. Signifi-cant differences between means were separated using Fisher’s Protected LSD at a=0.05.
3. Results
Seedlings with roots exposed to 10 and 20% ethanol solutions showed severe signs of needle burn within a week of treatment initiation. Nearly all seedlings in the 20% treatment and about half in the 10% treatment died within this time. Brown-ing of needles started at the base and moved toward the tip. Seedlings in the 0, 1, and 5% treatments showed no visible signs of damage during the first week. After (2 weeks) initiation, nearly 100% of the seedlings from the 20% treat-ment and approximately 90% of the seedlings in the 10% ethanol treatment were dead.
Ethanol was rapidly taken up by seedlings and concentrations in their needles and stems corre-sponded to the differences in ethanol treatments (Fig. 1). For each tissue, ethanol concentrations were significantly influenced by treatment solution and time since treatment initiation (PB0.001). In both 0 and 1% treatments, needle ethanol concen-trations were significantly high at 2 h (PB0.001), but by 4 h had dropped to their respective equi-librium levels. Needle ethanol concentrations from the 10% ethanol treatment continued to increase up to 8 h before leveling off. In contrast, stem ethanol concentrations increased to a maximum at 4 h for both the 1 and 10% ethanol solutions and then leveled off. Stem ethanol concentrations in control seedlings (0% ethanol) were near 0 and remained relatively unchanged over the course of 24 h.
Fig. 2. Ethanol concentrations in needles and stems of Dou-glas-fir seedlings (A) and in the surrounding air (B) when exposed to ethanol vapors from treatment applications. Points represent means 91 S.E. (n=3).
After 1 week (24 h after the third ethanol treat-ment), ethanol concentrations in needles (all P
values 50.016) and stems (all P values 50.013) continued to correspond to concentrations in the treatment solutions (Fig. 3). However, stem ethanol was several-fold higher than the needles for both the 1 and 5% ethanol treatments. There was no difference between ethanol concentrations at 07:30 and 15:30 h for needles or stems (data not shown). After 1 week of treatments, the midmorn-ing and afternoon net photosynthesis (Pn, Fig. 4A) was significantly different with 0\1\5% ethanol (all P values 50.033). Stomatal conductance (gs)
in the 0% treatment was significantly higher than either the 1 or 5% ethanol treatments at all times during the day (all P values 50.003) (Fig. 4C). Although gs in the 1% ethanol treatment was 1.5
times higher than the 5% treatment, it was not significantly different (P=0.139). Transpiration (E) differed among treatments with 0\1\5% throughout the day (all P values 50.001), except at 06:40 h whenEwas zero for all treatments (Fig. 4E). Afternoon E increased much more in the 0% treatments than in either the 1 or 5% treatments. Dark respiration at 22:30 h was 1.42mmol m−2 s−1 for all ethanol treatments.
At the end of 8 weeks (24 h after the final treatment), midmorning Pnwas still different (Fig. 4B) with 0\1\5% ethanol (all P values 5
0.024), although thePnof the 1 and 5% treatments
were close to zero. Stomatal conductance of con-trols was higher than either the 1 or 5% treatments (both Pvalues =0.002), both of which were close to zero (Fig. 4D). Transpiration by controls was higher than either the 1 or 5% treatments (both
P50.003), both of which were close to zero (Fig. 4F).
Xylem pressure potential 24 h after the initial ethanol treatment was lowest in the 5% treatment (P50.050), with no difference between the 1% treatment and controls (P=0.800) (Fig. 5). After 1 week, there were small differences in XPP among treatments with 0\1\5% ethanol (all
P50.049). After 8 weeks, the XPP again showed no difference between 0 and 1% treatments (P= 0.156), whereas it was significantly more negative in the 5% treatment (PB0.001 for comparisons with 0 and 1%) and most seedlings appeared dead (Fig. 5). Also at 8 weeks, shoot dry weights were higher for the controls than either the 1 or 5% treatments (both P values 50.026; Fig. 6A), and
there was a weak treatment effect on shoot lengths (P=0.082; Fig. 6B) with controls being the longest. There was no difference in these growth parameters between the 1 and 5% ethanol treatments.
4. Discussion
Ethanol solutions applied to roots at concentra-tions from 1 to 20% were injurious to Douglas-fir seedlings and did not enhance growth. Concentra-tions of 10 and 20% ethanol were lethal to seedlings within a week after initiating treatments and caused acropetal browning of needles. Ethanol concentrations above 10% applied to roots of tomatoes produced similar phytotoxic effects, and 5% concentrations severely reduced growth [11]. Douglas-fir seedlings were unable to tolerate 5% ethanol solutions. One week after initiating treatments, they had decreased net pho-tosynthesis, stomatal conductance, and water up-take (lower water potential) compared to controls, and after 8 weeks of treatments they were dying or dead. Seedlings treated with 1% ethanol showed limited damage at 8 weeks, but they may have eventually died because their gas exchange and biomass accumulation had declined significantly relative to the controls. If Douglas-fir seedlings have a threshold for tolerating ethanol applied externally to their roots, it probably occurs at a concentration of less than 1%.
Ethanol applied to roots of Douglas-fir seedlings was rapidly absorbed and transported up the stem where it accumulated to higher concen-trations than in needles. This is similar to the response observed when ethanol is synthesized in roots of flooded seedlings [5]. The ethanol does not accumulate in needles probably because a greater proportion of their cells are metabolically active and can metabolize ethanol into other or-ganic constituents [6 – 8]. Some ethanol may also be lost from the transpiration stream in needles and stems, but less than 5% of the ethanol fed to leaves of eastern cottonwood was transpired [8] and less than 0.2% of the ethanol generated in flooded roots of lodgepole pine escaped through stem lenticels [22]. Stem tissue may accumulate ethanol in part because of high levels of lignin which preferentially binds to alcohols [23]. The high ethanol concentrations in stem tissues may have damaged the cambium prior to any injury in the needles.
Applying 1% ethanol to the roots of Douglas-fir seedlings resulted in tissue concentrations ranging from 14.7 – 26.7 mmol g−1 fresh wt in stems and 1.4 – 2.3 mmol g−1 fresh wt in needles during the first week of treatments. These were 3 – 7 times
Fig. 4. Diurnal changes in net photosynthesis,Pn (A),
Fig. 5. Midmorning xylem pressure potential of Douglas-fir seedlings after 24 h, 1 week, and 8 weeks of treatment with 0, 1, or 5% ethanol solutions. Vertical bars are means 91 S.E. (n=3).
higher than the maximum ethanol concentrations in tissues of flooded Douglas-fir seedlings (stem 5.4 mmol g−1 fresh wt; needles 0.32 mmol g−1 fresh wt) [5]. Excised roots from Douglas-fir seedlings can quickly synthesize ethanol concen-trations from about 7.4 to 16.4 mmol g−1 fresh wt (31.1 – 76.8 mmol g−1 dry wt) when subjected to 4 h of N2 anoxia, depending on availability of
tissue nitrogen and soluble sugar concentrations [24]. But when the entire root systems of seedlings are flooded, a substantial portion of the ethanol synthesized in the roots diffuses into the surrounding water [5], decreasing the amount of ethanol moving upward in the transpirational stream. Although ethanol concentrations ]1% are injurious to Douglas-fir when applied through the roots, concentrations this high are unlikely to occur naturally, even under flooded conditions.
Douglas-fir needles readily absorbed ethanol from the atmosphere, while stems did not. Stom-atal pores may allow free diffusion of ethanol into the needles, whereas stem epidermal tissue is apparently impervious to atmospheric ethanol vapors. Rapid absorption of atmospheric ethanol by needles of control seedlings caused a tempo-rary (B2 h) increase in concentrations when control seedlings were positioned adjacent to the other ethanol treatments (Fig. 1A). We doubt that this transient spike of ethanol had any significant or lasting effect on the needles of control seedlings, but atmospheric cross-contamination needs to be considered when ap-plying ethanol treatments.
Applying ethanol to the roots of Douglas-fir seedlings at concentrations ]1% may have af-fected Pn directly in the needles or indirectly by
reducing gs. Ethanol inhibits enzyme synthesis in the aleurone layers of barley [10] and could
po-tentially inhibit critical photosynthetic enzymes in the chloroplast of conifers. Ethanol can also damage membrane integrity as observed in pea protoplasts at concentrations \0.92% [9]. Since thylakoid membranes play an important role in the light reactions of photosynthesis [18], any ef-fect of ethanol on this membrane could cause photosynthesis to drop, as observed in our seedlings.
Exposing roots of Douglas-fir seedlings to ethanol concentrations ]1% caused decreased gs
and transpiration rates at 1 and 8 weeks after treatments were initiated. In comparison, when Douglas-fir shoots were fed 0.1% ethanol there
was no effect on gs [5], indicating that processes
regulating gas exchange through stomata are af-fected by exposing roots to ethanol concentrations between 0.1 and 1.0%. Lower gs in seedlings treated with 1% ethanol did not result from de-creased water potentials, suggesting that ethanol may interfere with guard cell processes regulating stomatal opening rather than hydraulic processes. For instance, toxins produced by the fungus
Helminthosporium spp. inhibit stomatal opening by causing guard cell membrane damage and so-lute leakage [25]. Ethanol at ]1% concentration may cause similar guard cell damage. Alterna-tively, ethanol may be oxidized to acetaldehyde, which at low concentrations (0.005%) is relatively more toxic than ethanol to cell metabolism [26]. After a week of treatments, the XPP of seedlings receiving 5% ethanol was lower than seedlings treated with 0 or 1% ethanol, indicating that water uptake was probably impaired. The XPP did not differ between 0 and 1% treatments after one week, but all gas exchange parameters were significantly lower at 1%. Therefore, foliar membranes were probably damaged in the 1% ethanol treatments which reduced their photosyn-thesis and stomatal conductance as discussed above.
In summary, applying ethanol to roots at con-centrations ]1% negatively affected the physiol-ogy and growth of Douglas-fir seedlings. Ethanol concentrations of 10% or higher were lethal to seedlings after a week, whereas most seedlings at the 5% ethanol treatment died within 8 weeks. A decline in XPP at the 5% ethanol treatment indicates that water uptake also was reduced by possible damage to roots. The decline in Pn, and
gs with ethanol treatments ]1% may result from
damage to membranes involved in photosynthesis and stomatal function.
Acknowledgements
We thank K. Cameron, Champion Interna-tional Co., for generously supplying seedlings, Dr L. Ganio for advice on statistical analysis, Dr Bill Winner and Dr Bill Ferrell for critical reviews of the manuscript. The use of trade names is for information and convenience of the reader and does not constitute official endorsement or ap-proval by the USDA.
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