low concentrations 0.05 of ethanol [10]. How- ever, placing etiolated cuttings from mung bean
Vigna radiata L. 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 cm
3
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 – 450 mmol m
− 2
s
− 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, vv 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.
Ethanol concentrations in needles and stems were quantified at 2, 4, 8, and 24 h after the initial
treatment using two seedlings selected randomly from the 0, 1, and 10 ethanol treatments.
Seedlings from the 5 and 20 ethanol treatments were not included in this analysis. Old needles
were removed from the main stem and weighed 200 mg fresh weight into an autosampler vial
before sealing with a septum. New, expanding needles were not sampled or included in any mea-
surements because they varied in size and numbers among seedlings. In addition, a 2 cm stem segment
beginning at the root collar was cut, weighed, and sealed in a vial. Sealed vials were heated at 102°C
for 30 min to deactivate enzymes before analysis.
Ethanol analysis
was performed
with a
headspace autosampler Perkin Elmer HS40, Nor- walk, CT connected to a gas chromatograph
Hewlett-Packard 5890, Palo Alto, CA with a JW 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 cm
2
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
Fig. 1. Ethanol concentrations in the needles A and stems B of Douglas-fir seedlings during the first 24 h after treat-
ment with 0, 1, or 10 ethanol solutions. Points represent means 9 1 S.E. n = 3.
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 cm
3
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
