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Soil Biology & Biochemistry 39 (2007) 2414–2419
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Short communication
A method for linking in situ activities of hydrolytic enzymes to
associated organisms in forest soils
Shufu Donga, Denise Brooksb,!, Melanie D. Jonesa, Susan J. Graystonb
a
Biology and Physical Geography Unit, UBC Okanagan, 3333 University Way, Kelowna, BC, Canada V1V 1V7
Department of Forest Sciences, University of British Columbia, 2424 Main Mall, Vancouver, BC, Canada V6T 1Z4
b
Received 24 November 2006; received in revised form 5 March 2007; accepted 23 March 2007
Available online 4 May 2007
Abstract
A root window-based, enzyme-imprinted, membrane system has been modified to enable visualization of the activities of hydrolytic
enzymes (acid phosphatase, aminopeptidase, chitinase, and b-glucosidase) in situ in forest soils. The approach can be used to correlate
the distribution of enzyme activity with visible features such as roots, mycorrhizas, or mycelial mats. In addition, it enables accurate
spatial soil sampling for analysis of microbial communities associated with enzyme activities. The substrates are colorimetric conjugates
of napthol, where color develops instantly in the field, or fluorimetric conjugates of 4-methylumbelliferone, whose fluorescent products
are detected by a gel-documenting system. The method will allow important questions about the relationship between taxonomic and
functional diversity of soil microorganisms to be addressed and identification of enzyme activity hot-spots in soil.
r 2007 Elsevier Ltd. All rights reserved.
Keywords: Enzymes; Imprinting; Nutrient cycling; Roots; Root windows
1. Introduction
Enzymes have an obligatory role in catalyzing soil
nutrient transformations (Burns and Dick, 2002). Measurement of soil enzyme activities has, therefore, been
recommended as an extremely pertinent method for
measuring changes in soil quality (Dick, 1992), soil
recovery from disturbance or stress (Decker et al., 1999),
and as the most appropriate indicator of microbial
function (Caldwell, 2005). There are currently many wellutilized enzyme assays based on colorimetric and fluorimetric substrates that employ rapid microplate techniques,
as reviewed by Caldwell (2005). However, these assays all
involve soil sampling followed by lab analysis, inevitably
resulting in changes in enzyme activities (Tabatabai, 1994).
Thus these methods, like those that probe for DNA and
RNA of specific enzymes in soils (Kelly, 2003; Wellington
et al., 2003), reveal only potential, not actual, enzyme
activity in soils.
!Corresponding author. Tel.: +1 778 888 3464; fax: +1 604 822 8645.
E-mail address: [email protected] (D. Brooks).
0038-0717/$ - see front matter r 2007 Elsevier Ltd. All rights reserved.
doi:10.1016/j.soilbio.2007.03.030
In their recent review of nitrogen cycling, Schimel and
Bennett (2004) argue that soil processes will only be
understood if they are studied at much a finer scale than is
possible with conventional, destructive soil sampling. Here,
we report novel in situ methods to detect hotspots of C, N
and P cycling activity in the soil profile. The methods
modify and extend the field-based, root window approach
of Grierson and Comerford (2000).
2. Method development
Root
windows
(transparent
acrylic
panel
(77 cm ! 52 cm ! 0.6 cm) with a 30 cm ! 30 cm trap door
(Grierson and Comerford 2000)) were installed in a range
of Douglas-fir (Pseudotsuga menziesii) stands in the field
for 5 months prior to imprinting. A membrane of either
chromatography (Whatman, Cat No. 3030-861) or filter
paper (Whatman, Cat. No. 1001 055), treated with either a
mixture of substrate and colorimetric reagent or a
fluorimetric substrate, was placed directly on the soil
surface and enzyme activity detected by the appearance of
either colored or fluorescent products on the membrane.
Optimal duration of imprinting was 30 min for all enzymes
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S. Dong et al. / Soil Biology & Biochemistry 39 (2007) 2414–2419
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Fig. 1. (a) Color or fluorescence development, expressed as gray value, of imprints from assays for acid phosphatase, aminopeptidase, chitinase, and bglucosidase, with increasing time of contact with rhizobox soil (data from two runs for each enzyme). Note that increasing gray value represents higher
levels for the fluorescent products of chitinase and b-glucosidase; whereas higher levels of the colored products of phosphatase and aminopeptidase result
in lower gray values. (b) Fluorescence of 4-methylumbelliferone residue, expressed as gray scale, adhering to untreated pieces of filter paper applied to
rootboxes containing Douglas-fir seedlings 1–8 days after initial assays (day 0) for chitinase and b-glucosidase activity. As fluorescence increases, gray
value increases. Mean values7SEM of three replicate rhizoboxes containing 6-months old. Douglas-fir seedlings growing in field soil.
except phosphatase, which required 60 min (Fig. 1a). After
exposure, the imprints were carefully removed, rinsed with
deionized water, air-dried and scanned. Membranes were
handled throughout with latex gloves or sterilized forceps.
To detect acid phosphatase activity, chromatography
paper was soaked for 1 min in a 1:10 (v/v) mixture of
freshly prepared 50 mM a-naphthyl phosphate (Sigma
N7255) and 10 mM Fast Red TR (Sigma F2768), both
prepared in 50 mM pH 5.6 citrate buffer (Dinkelaker and
Marschner 1992), and then air-dried. Standards (Sigma
P3627, from wheat germ) of 0–0.35 enzyme units
(EU) ml"1 in 5 ml pH 5.6 citrate buffer were applied to
separate pieces of membrane and placed adjacent to test
membranes on soil surfaces The intensity of purple-red
color after conversion to gray scale in Adobe Photoshop
Elements 2.0, represented acid phosphatase activity
(Fig. 2a). Control membranes treated with only Fast Red
showed no color after imprinting.
Membranes to detect aminopeptidase activity were
prepared by soaking in 20 mM L-leucyl 2-naphthylamide
(Sigma L0376, prepared in 95% alcohol) followed by airdrying. Fast Blue BB (2.4 mM in DI water, Sigma F0250)
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S. Dong et al. / Soil Biology & Biochemistry 39 (2007) 2414–2419
Fig. 2. Imprints and soil profiles from root windows in interior Douglas-fir stands near Barriere, British Columbia, Canada. (a) phosphatase imprint; (b)
aminopeptidase imprint, (c) soil image overlain with the same imprint, (d) image of soil profile; (f) b-glucosidase imprint and (e)associated soil image; (h)
chitinase imprint and (g) associated soil image. All images are at the same scale.
was applied in a fine mist after imprints and standards were
removed from soil surfaces. Imprints were then exposed to
150 W infrared light (1 min) to minimize development of
non-specific background color (Humble et al., 1977). An
orange-red color represented aminopeptidase activity.
Standards for the aminopeptidase assay were prepared
from a fungal protease/peptidase complex of Aspergillus
oryzae (Sigma, P6110) and applied to membranes in 5 ml
aliquots containing 0, 3.9, 7.8, 15.6, and 31.2 EU ml"1.
Control membranes received no substrate and exhibited no
color when sprayed with Fast Blue. Although the resolution of aminopeptidase activity was lower than acid
phosphatase, the association of aminopeptidase activity
with roots could be clearly observed (Fig. 2b–d).
Chitinase activity was visualized on membranes soaked
in 5 mM 4-methylumbelliferyl-N-acetyl-b-glucosaminide
(Sigma M2133) in 2-methoxylethanol (Sigma M5378).
Membranes for b-glucosidase used 4-methylumbelliferylb-glucopyranoside dehydrate (Sigma M3633) as a substrate. Activity of these enzymes on these substrates release
fluorescent 4-MUB (Hoppe, 1983; Pritsch et al., 2004). In
the lab imprints were imaged with a gel documentation
system (Gel LOGIC 440, Eastman Kodak Company, New
Haven, CT, USA). Persistence of fluorescent residues on
soil surfaces was found for 8 days following imprinting
(Fig. 1b), so we recommend that tests using 4-MUB-linked
substrates be at least 10 days apart. 4-MUB (Sigma
M1381) standards of 0, 0.078, 0.156, 0.313, and
0.625 mM in 2-methoxylethanol were applied in 5 ml
aliquots to membranes to generate standard curves for
both chitinase and b-glucosidase activity. When test
membranes prepared with either the substrate for chitinase
or the substrate for b-glucosidase were applied to soil
surfaces, fluorescence could be detected on the resulting
imprints (Fig. 2e–h). The boundaries of the fluorescent
spots were diffuse but activities could be associated with
specific structures in the soil. Control membranes exhibited
little fluorescence, with gray values typically below 100.
3. Applications of the method
By generating standard curves either from enzyme
standards applied to test membranes in the field (phosphatase and aminopeptidase) or from known amount of
product applied to membranes just before imaging (MU
for chitinase and b-glucosidase), it is possible to use the
method to semi-quantitatively estimate in situ enzyme
activities. For example, imprints taken at root windows
installed in a mixed Picea engelmannii/Abies lasiocarpa
stand (Hagerman et al., 1999) detected significantly higher
phosphatase activities in nine natural stands (24.472.1)
than in nine clear cuts (6.572.2). The method also allows
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Fig. 3. Phosphatase activity derived from scanned 20 cm ! 20 cm imprints taken from a Douglas-fir/birch chronosequence of four ages: young (3–6 years),
canopy closure (24–27 years), stem exclusion (55–60 years), and older (88–100 years) near Enderby, British Columbia, Canada (mean values7SEM for
total number of active areas; n ¼ 3). Different letters indicate a significant difference at Po0.05 according to Tukey test for the total number of active
areas illustrated in the bar. (a) Total number of visible phosphatase-active areas per imprint. (b) Frequency distribution of different sizes of phosphataseactive areas larger than 50 pixels in size on scanned imprints.
determination of the extent and spatial distribution of
enzyme activity. For example, the observation that the
number, average size, and intensity of phosphatase hot
spots increased with stand age in mixed P. menziesii/Betula
papyrifera stands (Fig. 3a and b), prompted us to
determine whether the composition of the microbial
community associated with high phosphatase patches
changes with stand age.
A major strength of the in situ methods described
here is that they can be used to correlate enzyme activities
with plant, fungal, and bacterial communities in soils prior
to destructive sampling. The location of macroscopic
features, such as soil horizons, roots, and mycelial fans,
can be recorded with high-resolution digital photography.
These images can then be overlain with scans of the
imprints, and the two images aligned using holes in the
imprints created by pins inserted through the root window
(Grierson and Comerford, 2000). Such an approach
has demonstrated that, in clear cut soils, b-glucosidase
and chitinase activities were found almost exclusively
in association with roots and decaying material,
whereas in adjacent Douglas-fir stands, activity was
mainly associated with fungal mats or organic horizons
(Fig. 4a and b).
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Fig. 4. Percentage of chitinase and b-glucosidase activity associated with organic soil (OS), mineral soil (MS), roots (RT), fungal mats (FM), and decayed
material (DM) on imprints taken in three root windows per site in (a) Douglas-fir forests and (b) clear cuts near Barriere, British Columbia, Canada (mean
values7SEM; n ¼ 4 sites).
By using molecular techniques such as DGGE or
T-RFLP, soil fungal and prokaryotic communities associated with hot spots of enzyme activity can be described.
To do this, the imprints can be used to create templates
from clear acetate sheets for sampling soils at mm scales.
This approach will complement approaches that probe for
specific genes (Kelly, 2003; Wellington et al., 2003) by
correlating the presence of organisms with directly
measured activity.
Many factors that affect enzyme activity, such as pH,
moisture and temperature, will vary at a fine scale across
individual root windows. Although this variation might be
seen as a disadvantage by researchers used to measuring
bulk soil enzyme activities, we see it as a major advantage
of the method. This method gives us confidence that we are
detecting actual enzyme activity as it occurs in the field in
soil microsites. This fine-scale variation in environmental
conditions is lost during typical soil sampling. Thus,
this is exactly the type of approach that will allow us to
study soils at the scales suggested by Schimel and Bennett
(2004) and hence, deepen our understanding of soil
processes.
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Acknowledgements
The authors gratefully acknowledge financial support
from the Special Research Opportunity and Discovery
Grant programs of the Natural Sciences and Engineering
Research Council of Canada, the Forest Science Program
of the Forest Investment Account of British Columbia, and
the award of a University of British Columbia Cordula and
Gunter Paetzold Fellowship to DB. We are grateful for
excellent field assistance from Jason Barker, Anne Bernhardt, Ben Chester, Julie Deslippe, Alanna Leverrier,
Kristen Mackay, Adam McCaffrey, Chelsea Ricketts,
and Jeff Sherstobitoff. This research was based on initial
discussions and considerable advice from Pauline Grierson
during a sabbatical leave by MJ at the University of
Western Australia.
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