Nutrient limitations to soil microbial biomass

and activity in loblolly pine forests

A.S. Allena, W.H. Schlesingera,b,*

aDepartment of Biology, Duke University, P.O. Box 90338, Durham, NC 27708-0338, USAbNicholas School of the Environment and Earth Sciences, Duke University, P.O. Box 90339, Durham, NC 27708-0329, USA

Received 15 August 2001; received in revised form 21 October 2003; accepted 17 December 2003

Abstract

We performed an assay of nutrient limitations to soil microbial biomass in forest floor material and intact cores of mineral soil collected

from three North Carolina loblolly pine (Pinus taeda) forests. We added solutions containing C, N or P alone and in all possible

combinations, and we measured the effects of these treatments on microbial biomass and on microbial respiration, which served as a proxy

for microbial activity, during a 7-day laboratory incubation at 22 8C. The C solution used was intended to simulate the initial products of fine

root decay. Additions of C dramatically increased respiration in both mineral soil and forest floor material, and C addition increased

microbial biomass C in the mineral soil. Additions of N increased respiration in forest floor material and increased microbial biomass N in the

mineral soil. Addition of P caused a small increase in forest floor respiration, but had no effect on microbial biomass.

q 2004 Elsevier Ltd. All rights reserved.

Keywords: Soil microorganisms; Nutrient limitation; Respiration; Nitrogen; Phosphorus; Carbon

1. Introduction

Soil microbial biomass plays a critical role in nutrient

retention and soil fertility in terrestrial ecosystems.

Microbial activity and biomass are tightly linked to N

mineralization in soils (Myrold, 1987; Hart et al., 1994;

Tietema, 1998). In addition, microbial respiration represents

the primary mechanism for degradation of carbon fixed by

plants, and microbial respiration rates may determine the

potential for C sequestration in the terrestrial biosphere

(Hungate et al., 1997; Schlesinger, 1997). Rising atmos-

pheric CO2 concentrations and atmospheric N deposition

are two important global change processes that may affect

soil microbes. If elevated CO2 increases inputs of organic C

to the soil by stimulating photosynthesis (Curtis and Wang,

1998), increased N mineralization by carbon-limited soil

microbes may increase soil N availability (Zak et al., 1993).

On the other hand, Dıaz et al. (1993) suggested that elevated

CO2 would cause C-limited microbes to grow rapidly

and outcompete plants for available nitrogen. If soil

microbes are limited by N, they may accumulate N

deposited from the atmosphere and buffer ecosystems

against harmful effects of N saturation (Aber et al., 1998).

Atmospheric N deposition may also cause N-limited soil

microbes to decompose organic matter more quickly and

reduce C storage in soil. Determining which resources limit

microbial biomass and activity will elucidate controls on

nutrient and carbon cycling in soils.

Microbial biomass appears to be closely linked to

aboveground plant productivity in many ecosystems (Zak

et al., 1994), suggesting that the biomass of microbes

depends directly on inputs of reduced carbon to the soil.

Direct additions of readily available carbon sources such as

glucose or sucrose to soil usually result in increases in

microbial activity or biomass (Anderson and Domsch, 1985;

Gallardo and Schlesinger, 1994; Jonasson et al., 1996a,b).

Microbial biomass and activity may also be limited by the

availability of N or P (Wardle, 1992). Gallardo and

Schlesinger (1994) found that addition of NH4NO3

increased microbial biomass N in the forest floor of a

warm-temperate hardwood forest. Immobilization of N by

microbial biomass plays a crucial role in ecosystem nutrient

retention in many forests, and tracer experiments using 15N

0038-0717/$ - see front matter q 2004 Elsevier Ltd. All rights reserved.

doi:10.1016/j.soilbio.2003.12.002

Soil Biology & Biochemistry 36 (2004) 581–589

www.elsevier.com/locate/soilbio

* Corresponding author. Address: Nicholas School of the Environment

and Earth Sciences, P.O. Box 90329, Duke University, Durham, NC 27708-

0329, USA. Tel.: þ1-919-613-8004; fax: þ1-919-613-8077.

E-mail address: schlesin@duke.edu (W.H. Schlesinger).

show that a large proportion of N added as 15NH4þ or 15NO3

2

moves quickly into microbial biomass (Vitousek and

Matson, 1984; Zak et al., 1990). The concentration of N

in forest floor material may be positively (Melillo et al.,

1982) or negatively (Magill and Aber, 1998) related to

microbial respiration, with negative effects prevailing in

forest floor material that has decomposed for several years

(Berg, 1986; Magill and Aber, 1998). Phosphorus limitation

to microbial biomass has been demonstrated in the mineral

soil of at least two temperate forests (Scheu, 1990; Gallardo

and Schlesinger, 1994). Gallardo and Schlesinger (1994)

suggested that microbial P limitation may be common in

highly weathered soils in which P tends to be bound in iron

or aluminum sesquioxides.

Loblolly pine (Pinus taeda) forests cover large areas in

the southeastern US, where they often develop rapidly on

abandoned agricultural land. Loblolly pines may respond to

rising atmospheric CO2 with greater photosynthetic rates

(Ellsworth, 1999), aboveground growth (DeLucia et al.,

1999), litterfall (Finzi et al., 2001), and root production

(Matamala and Schlesinger, 2000). Leaching of organic C

compounds from leaves can also be accelerated by elevated

CO2 (Lichter et al., 2000). Understanding how soil microbes

respond to increased C, N and P availability may add to our

understanding of the mechanisms by which elevated CO2

and N deposition affects ecosystem function in loblolly pine

forests.

In this study, we examine the effects of C, N and P

additions on microbial biomass C and N and microbial

respiration in forest floor samples and in intact mineral soil

cores taken from three loblolly pine stands in central North

Carolina.

2. Methods

2.1. Experimental design and sampling

We selected three loblolly pine stands, each 16 years old,

in the Duke Forest near Durham, North Carolina. In order to

include a range of soils, we selected sites differing in soil

series, pH and texture (Table 1). At each site, we selected

a study area measuring 30 £ 40 m2, in which we randomly

selected four points. Each point acted as an experimental

block, from which forest floor material was sampled within

a 30 £ 30 cm2 plot. After removing the forest floor, we

collected eight cores of mineral soil in each plot, with each

core measuring 4.7 cm in diameter and 7.5 cm in depth

(about 105 g dry weight). We sampled two of the four

blocks in each stand during June 1998 and the two

remaining blocks during November 1998.

We collected soil cores using a slide-hammer coring

device (AMS, Inc.; American Falls, ID) that held 8.5-cm

long removable plastic sleeves with an inside diameter of

4.7 cm. Each core was left in its sleeve throughout the

experiment to preserve soil structure and prevent mixing of

soil that might accelerate microbial activity. After sampling,

we placed cores in an ice chest until returning to the

laboratory (within 8 h). Cores were then stored at 4 8C

overnight before beginning the fertilization and incubation

procedures the following day.

We used scissors to cut large pieces of forest floor

samples into pieces #5 cm to facilitate homogenization and

subsampling within blocks. For each forest floor sample

(representing a single block), we placed subsamples of

approximately 9.3-g dry weight into each of eight 1 -l

mason jars with lids that had been fitted with butyl rubber

septa.

2.2. Nutrient solution application

We pipetted 5 ml of one of the eight nutrient solutions

(reflecting the eight possible combinations of C, N and P

treatments or de-ionized H2O, Table 2) onto each forest

floor subsample and mixed the material for several seconds

with a spatula to distribute the nutrient solution as evenly as

possible. This resulted in additions of 27 000 mg C g21,

540 mg N g21 and 54 mg P g21 to the forest floor samples.

Control samples received 5 ml of de-ionized H2O. The C

solution used was intended to simulate initial products of

fine root decay (J.M. Stark and S.C Hart, pers. comm.). We

injected each soil core with 7 ml of one of the eight nutrient

solutions using a modified spinal needle to minimize

disturbance to soil structure. To create side-port needles,

Table 1

Site characteristics of the three 16-year-old loblolly pine stands used in this study

Site 1 Site 2 Site 3a

Soil taxonomy Typic Kanhapludult (Appling series) Aquic Hapludult (Helena series) Ultic Hapludalf (Enon series)

pHCaCl2

b 4.6 3.5 5.0

Texture Loam Sandy loam Sandy loam

Site history Loblolly and shortleaf pines established from

about 1900 to 1935; cut (except seed trees)

1982; disked 1982; seed trees removed 1985

Loblolly and shortleaf pines established

about 1900; cut (except seed trees) 1982;

burned 1982

Loblolly pines established during 1930s

and 1940s; cut 1983; drum chopped and

burned 1983; loblolly pines planted 1983

a Site 3 is the same loblolly pine stand used in the Duke Forest free-air CO2 enrichment (FACE) experiment. Samples for the present study were taken at least

40 m away from any FACE experimental plot.b pH in 0.01 M CaCl2 is typically lower than pH in water by about 0.5 pH units.

A.S. Allen, W.H. Schlesinger / Soil Biology & Biochemistry 36 (2004) 581–589582

we used a sharp-edged file to make two small holes on

opposite sides of 3.5 in., 18-gauge needles (Becton Dick-

inson and Co., Franklin Lakes, NJ, item #405184), about 6

and 8 mm from the tip. We plugged the tip of each needle by

cutting the last 5 mm from the obdurator, and we glued this

short wire segment into the tip of the needle using a

cyanoacrylate glue. When inserting the needle into a soil

core, we plugged the inside of the needle with the remaining

long segment of the obdurator to prevent clogging with soil.

We then removed the obdurator and attached a 1-cm3

tuberculin syringe that had been filled with the appropriate

nutrient solution. We slowly pressed the syringe plunger

while withdrawing the needle to distribute the solution as

evenly as possible through the length of the core. This

procedure was repeated a total of seven times for each core

at evenly spaced injection points. A small amount of

nutrient solution leaked from the bottom of some cores

during the injection procedure, but the amounts actually

applied to cores were within 10% of the 7-ml target

quantity. Immediately after injection, cores were placed in

mason jars as described above.

The nutrient solutions added to the mineral soil samples

resulted in additions of 3300 mg C g21, 66 mg N g21 and

6.6 mg P g21 to each core. Control samples received 7 ml of

de-ionized H2O. This C addition rate to mineral soil,

combined with the C addition rate to forest floor material,

provided a supplement of approximately 260 g C m22,

which is similar to the rate of C deposition measured in

litterfall at the Duke Forest FACE site (site 3 in this study;

DeLucia et al., 1999). Ratios of C:N and C:P in added

solutions were similar to the ratios in green leaf tissue.

2.3. Respiration

Immediately after applying nutrient solution to a soil

core or forest floor sample, we ventilated its mason jar with

room air (usually about 400 ml l21 CO2) for several seconds

using a hand-held electric fan, and then closed the jar

tightly. We measured concentrations of CO2 in air using an

infrared gas analyzer (EGM-1; PP Systems, Inc.; Haverhill,

MA). After approximately 4 h, we used a syringe to remove

a 10-ml sample from the headspace of each jar through its

septum and injected this sample into the infrared gas

analyzer for CO2 concentration measurement.

Between measurement periods, jars were covered loosely

with perforated aluminum foil and stored in a dark cabinet at

approximately 22 8C. Water content was maintained at a

level equal to field moisture plus the fertilizer or H2O

solution by weighing the jars after adding treatment

solutions and adding de-ionized water with a spray bottle

several times during the 7-day period. Jars were vented and

closed for measurements of CO2 accumulation six times

during the following week. Jars were left closed to allow

accumulation of CO2 for approximately 4 h on the first few

days, and incubation times were lengthened to approxi-

mately 7 h during the last days of the incubation when

respiration rates had declined. Times were chosen to be long

enough to allow sufficient CO2 accumulation for accurate

measurement in all samples (i.e. .0.15% [CO2]), but short

enough to avoid excessive CO2 concentrations that may

cause negative feedbacks on microbial respiration (i.e.

.4% CO2; Sierra and Renault, 1995). Concentrations of

CO2 higher than 2% were generally avoided, although in a

few cases, samples with high respiration rates exceeded 3%

CO2. Because some air with above-ambient CO2 may have

remained inside soil cores during the flushing procedure,

the respiration rates measured here may overestimate

actual rates.

2.4. Microbial biomass C and N

After the 7-day incubation period, soil cores were

removed from their sleeves, passed through a 5-mm sieve,

and subsampled for measurement of microbial biomass C

and N using a fumigation–extraction procedure (Brookes

et al., 1985; Vance et al., 1987; Gallardo and Schlesinger,

1990). Briefly, two subsamples of 8 g of mineral soil or

1.5 g of forest floor material (dry weights) were weighed

into 50-ml centrifuge tubes. One subsample was extracted

immediately with 40 ml of 0.5 M K2SO4 solution, shaken

for 1 h on an oscillating shaker, and filtered with a Whatman

No. 1 filter that had previously been rinsed with K2SO4

followed by de-ionized water. The second subsample was

fumigated with 3 ml of ethanol-free chloroform that was

pipetted onto two large cotton balls that were placed in the

headspace of the centrifuge tube. Each fumigated tube was

tightly capped immediately after adding chloroform and

stored in a dark cabinet for 7 days. A preliminary test of this

fumigation procedure gave results identical to those

obtained when samples were fumigated in a desiccator as

described by Jenkinson and Powlson (1976). To remove

chloroform at the end of this fumigation period, we opened

the tubes, removed cotton balls, and placed the tubes in a

large glass vacuum desiccator. We evacuated the desiccator

eight times for 3 min each, flushing the desiccator with

room air after each evacuation (Horwath and Paul, 1994).

These tubes were then extracted with K2SO4 as described

above. We measured N in the extracts using a persulfate

Table 2

Ingredients of the treatment solutions

Treatment Ingredient Concentration

C Cellobiose (15 g C l21)

Vanillic acid (10 g C l21)

Pectin (10 g C l21)

Na citrate (5 g C l21)

Sucrose (5 g C l21)

Mannose (5 g C l21)

N NH4NO3 (1 g N l21)

P KH2PO4 (0.1 g P l21)

The C, N and P treatments were used alone or in combination with other

treatments. Treatment solutions with C contained a total of 50 g C l21.

A.S. Allen, W.H. Schlesinger / Soil Biology & Biochemistry 36 (2004) 581–589 583

oxidation procedure (D’Elia et al., 1977) followed by

colorimetric NO32 analysis (TRAACS 800 Autoanalyzer,

Bran Leubbe, Elmsford, NY). Preliminary tests showed that

this method recovered .90% of organic N in glutamic

acid standards. We measured C in extracts using a

Shimadzu TOC 5000 solution C analyzer (Shimadzu, Inc.,

Columbia, MD).

2.5. Statistical analyses

In order to estimate the total quantity of CO2 evolved

from each sample during the 7-day incubation period, we

calculated the areas under straight lines drawn between the

respiration rate data points. We analyzed data using a nested

Analysis of Variance (ANOVA) design (DataDesk, Data

Description Institute, Ithaca, NY), with blocks acting as a

random effect nested within ‘Date’ (i.e. June and Novem-

ber) and ‘Site.’ ANOVA tables for forest floor respiration,

mineral soil respiration and microbial biomass are found in

Allen (1999). The error terms and denominator degrees of

freedom for treatment effects are derived from the

interaction between ‘block’ and the treatment of interest.

Respiration data were log transformed prior to ANOVA to

homogenize variance.

Treatment means were compared using Scheffe’s post

hoc test. For parameters in which we found interactions

between sampling date and other factors (i.e. forest floor

respiration and forest floor microbial biomass N), we

calculated Scheffe-Test P values for comparisons within

dates, and we created a separate graph for each date. For

other parameters, we calculated Scheffe-Test P values

comparing groups that combine data from the two dates and

three sites.

3. Results

3.1. Forest floor

Addition of organic C caused a dramatic stimulation of

respiration rates in forest floor samples in June and

November (P , 0:0001 on both sampling dates, Scheffe’s

Tests). Respiration peaked 1 day after injecting C solutions

(Fig. 1A and B), when C-treated samples exhibited

respiration rates that were about four times higher than the

rates seen in the H2O-treated control samples. C-treated

samples respired an average of 21 756 mg C g21 forest floor

by the end of the 7-day incubation period, which is about

three times larger than the total respiration from samples

that received only H2O (P , 0:0001 in June and November,

Scheffe’s Tests). The effect of C addition on cumulative

respiration over the 7-day incubation period in forest floor

samples varied between the two sampling dates

(P ¼ 0:0042 for ANOVA Date by C interaction), largely

because samples that did not receive C respired faster in

November than in June (P , 0:0001; Scheffe’s Test).

Addition of N to forest floor samples caused a small but

significant increase in respiration rate on days 2–7 of the

incubation (P , 0:02 in June and November, Scheffe’s

Tests; Fig. 1A and B). After 7 days of incubation, N-treated

forest floor samples had respired 30% more C than control

samples (P , 0:001 in June and November, Scheffe’s

Tests). The effect of N on cumulative respiration in forest

floor samples after 7 days was larger in November than in

June (Date by N interaction, P ¼ 0:0200; ANOVA),

although the effect of N was significant at both sampling

times (P ¼ 0:0082 in June and P ¼ 0:0002 in November,

Scheffe’s Test). Addition of N also interacted with C

addition on the second day of the incubation period,

resulting in significantly higher respiration in C þ N-treated

samples as compared with C-treated samples (P , 0:001 in

June and November, Scheffe’s Tests). However, respiration

declined more rapidly in C þ N-treated samples than in C-

only samples, and after 3 days of incubation, C þ N-treated

samples respired at lower rates than C-treated samples

(P , 0:03 in June and November, Scheffe’s Test). The

initial, positive interaction between C and N resulted in a

slight increase of more than 10% in total CO2 respired after

7 days in C þ N-treated forest floor samples as compared

with C-treated forest floor samples (P , 0:03 in June and

November, Scheffe’s Test).

Phosphorus addition caused a small but statistically

significant increase in respiration of forest floor samples that

received no other treatment (P , 0:03 in June and

November, Scheffe’s Test; Fig. 1A and B), but P addition

did not increase respiration rates in samples that also

received C, N or C þ N treatments.

Addition of C to forest floor material had no significant

effect on microbial biomass C or N (P . 0:3 in all cases,

Scheffe’s Test; Figs. 2 and 3). Addition of N to forest floor

material did not significantly increase microbial biomass C

when N was added alone or in conjunction with other

treatments (P . 0:05 for ANOVA main effect of N and

interactions with N; Fig. 2). However, microbial biomass N

in forest floor increased when N was added in the absence of

other treatments in November (P ¼ 0:008; Scheffe’s Test;

Fig. 3B). Addition of N alone did not significantly affect

forest floor microbial biomass N in June (P ¼ 0:502;

Scheffe’s Test; Fig. 3A).

3.2. Mineral soil

Addition of organic C to the mineral soil caused

significant, order-of-magnitude increases in soil respiration

rates within hours of C addition (Fig. 4). Respiration rates in

mineral soil samples that received C peaked 2 days after C

fertilization, when the mean respiration rate in these

samples was six times higher than the mean rate in samples

that received only H2O (P , 0:0001; Scheffe’s Test). Seven

days after nutrient additions, the mean respiration rate in

mineral soil samples that received C was still four times

higher than in samples that received only H2O (P , 0:0001;

A.S. Allen, W.H. Schlesinger / Soil Biology & Biochemistry 36 (2004) 581–589584

Scheffe’s Test). The total quantity of C respired by C-treated

mineral soil during the 7-day incubation period was

1223 mg C g21 soil, which is about six times greater than

the total C respired by mineral soil samples that received

only H2O (P , 0:0001; Scheffe’s Test).

Mineral soil samples that received N alone had higher

microbial respiration than control samples, but this

difference was not statistically significant (P . 0:10 on all

dates, Scheffe’s Test; Fig. 4). On day 2 of the incubation, a

positive interaction between C and N appeared, such that

C þ N-treated mineral soil had nearly double the respiration

rates observed in C-treated mineral soil (P ¼ 0:0214;

Scheffe’s Test). This interaction disappeared completely

after 4 days of incubation, and after 7 days, the interaction

reversed so that C þ N-treated mineral soil had lower

respiration rates than C-treated mineral soil (P ¼ 0:0418;

Scheffe’s Test). The total quantity of C respired by C þ N-

treated mineral soil after 7 days of incubation was not

significantly different from that respired by ‘C-only’

samples (P ¼ 0:1099; Scheffe’s Test). Addition of P did

not significantly affect respiration rates in the mineral soil

(P . 0:10 on all dates, ANOVA; Fig. 4). Cumulative

respiration in ‘control’ mineral soil samples did not

differ significantly among sites (P . 0:05; Sheffe’s Tests;

Table 3).

After 1 week of incubation, C addition increased

microbial biomass C in the mineral soil by 40%

(P ¼ 0:0042; Scheffe’s Test; Fig. 5). However, addition of

C alone did not significantly increase microbial biomass N in

the mineral soil (P ¼ 0:2765; Scheffe’s Test; Fig. 6).

Addition of N alone to the mineral soil significantly increased

microbial biomass N (P ¼ 0:016; Scheffe’s Test; Fig. 6),

Fig. 1. Microbial respiration in forest floor material taken from three loblolly pine stands in central North Carolina in June, 1998 (A) or November, 1998 (B),

incubated in mason jars in the laboratory at 22 8C. Forest floor subsamples received injections of C, N or P solutions, alone or in combination, just prior to the

first respiration measurement (P treatment data not shown). Respiration rates were measured during CO2-measurement periods of several hours when jars were

sealed. Data shown reflect averages across the three sites. Scatter of points in X direction (i.e. around sampling times) is exaggerated for clarity. Points on the

same day with the same letter are not significantly different (P . 0:05; Scheffe Test). Error bars are 1 standard error, calculated using only the six values for a

given treatment. These error bars include some variability due to differences among sites that is accounted for in the ANOVA used to produce the Scheffe

statistics.

A.S. Allen, W.H. Schlesinger / Soil Biology & Biochemistry 36 (2004) 581–589 585

but addition of N increased microbial biomass C only when N

was added in conjunction with C (P ¼ 0:0071 for ANOVA C

by N interaction; Fig. 5).

The ratio of C to N in microbial biomass in mineral soil

was altered by a three-way, Date-by-C-by-N interaction

(P , 0:0001; ANOVA). C addition increased the microbial

biomass C:N ratio in the presence and absence of N on each

date (P , 0:0001 in each case, Scheffe’s Tests). The ratio of

C:N in microbial biomass was consistently lower with

nitrogen addition. This difference was statistically signifi-

cant in the presence and absence of C in June 1998, and in

the presence of C additions in November 1998 (P , 0:001

in each case, Scheffe’s Tests).

4. Discussion

We expected to find a C limitation of microbial activity

in the mineral soil and limitation by N or P in forest floor

material. Microbial respiration in the mineral soil and in

forest floor increased dramatically with addition of labile C

(Figs. 1 and 4), while additions of N and P did not

significantly alter respiration rates in the mineral soil and

they had only small effects on respiration in forest floor

material. Our results suggest that if the concentration of

Fig. 2. Microbial biomass C in forest floor material from three loblolly pine

stands in central North Carolina, 7 days after applying C or N solutions

alone or in combination (P treatments not shown). Data are averages of

values from June and November sampling dates. Error bars are 1 standard

error, calculated using only the eight values for a given treatment. These

error bars include some variability due to differences among sites and dates

that is accounted for in the ANOVA used to produce the Scheffe statistics.

Microbial biomass C was determined by chloroform fumigation–extrac-

tion, and dissolved organic C values (determined by catalyzed combustion)

were divided by a KEC correction coefficient of 0.45 to convert to biomass

values. Bars with the same letter are not significantly different (P . 0:05;

Scheffe’s Test).

Fig. 3. Microbial biomass N in forest floor material collected in June, 1998

(A) and November 1998 (B) from three loblolly pine stands in central North

Carolina, 7 days after applying C or N solutions alone or in combination (P

treatments not shown). Microbial biomass N was determined by chloroform

fumigation–extraction, and dissolved organic N values (determined by

persulfate digestion and automated NO32 analysis) were divided by a KEN

correction coefficient of 0.54 to convert to biomass values. Error bars are 1

standard error, calculated using only the six values for a given treatment.

These error bars include some variability due to differences among sites that

is accounted for in the ANOVA used to produce the Scheffe statistics. Bars

with the same letter on a given day are not significantly different (P . 0:05;

Scheffe’s Test).

Fig. 4. Microbial respiration in intact cores of mineral soil (0–7.5 cm

depth) taken from three loblolly pine stands in central North Carolina,

incubated in mason jars in the laboratory at 22 8C. Data shown reflect

averages across two dates (June and November 1998) and the three sites.

Cores received injections of C, N or P solutions, alone or in combination,

just prior to the first respiration measurement. Scatter of points in X

direction (i.e. around sampling times) is exaggerated for clarity. Points on

the same date with the same letter are not significantly different (P . 0:05;

Scheffe Test). Error bars are 1 standard error, calculated using only the 12

values for a given treatment. These error bars include some variability due

to differences among sites and dates that is accounted for in the ANOVA

used to produce the Scheffe statistics. (A) Respiration rates measured

during CO2-measurement periods of several hours when jar was sealed. (B)

Cumulative respiration, estimated by calculating the area under straight

lines drawn between rate data points.

A.S. Allen, W.H. Schlesinger / Soil Biology & Biochemistry 36 (2004) 581–589586

relatively labile C in plant litter increases due to a

perturbation such as rising atmospheric CO2, decomposition

rates could increase.

We found that cumulative respiration in mineral soil

samples fertilized with C alone exceeded respiration in

control samples by an average of 1009 mg C g21 soil, an

amount equal to about 30% of the C added. Cumulative

respiration in forest floor samples that received only C

exceeded respiration in samples that received H2O by an

average of 14 764 mg C g21 soil, a quantity of C equal to

55% of the C added. This rapid consumption of the added C

may explain the convergence of respiration rates between

C-fertilized and non-C fertilized mineral soil and forest floor

samples by the end of the incubation period.

Melillo et al. (1982) suggest that microbial decompo-

sition of leaf litter should increase with increasing N

availability to microbes. In the present study, addition of N

to forest floor samples increased respiration rates by an

average of 30% over the 7-day incubation period, and the

percentage increase in decomposition rate due to N addition

increased during the incubation period from 19% on the first

day to 37% after 7 days (Fig. 1). Although, the

concentration of NH4NO3 added in the present experiment

(1 g N l21) was about three orders of magnitude greater than

typical rainfall concentrations of NH4þ and NO3

2 in this

region (NADP, 1999), the strong and consistent respiration

response suggests there is a potential for accelerated loss of

forest floor material if atmospheric N deposition increases in

the future. In central North Carolina, the greatest litterfall

occurs during October, and our November collection

contained a larger proportion of recently senesced litter

than our June collection. Interestingly, we found that

respiration in forest floor material responded more strongly

to addition of N in November than in June (Fig. 1). Berg

(1986) suggested that decomposition of Pinus sylvestris

litter is initially limited by availability of N and P as

cellulose is decomposed, but subsequent decomposition of

lignin is retarded by N additions. Our results are consistent

with this model, insofar as the forest floor material collected

in November was dominated by cellulose. Microbial

biomass N in both forest floor and mineral soil increased

significantly when N was added alone in November

(Fig. 3B), suggesting these pools could accumulate N

derived from increasing atmospheric N deposition.

Microbial biomass C in the mineral soil increased when

C was added (Fig. 4). This result is consistent with the

prediction of C limitation to microbes in the mineral soil. It

is also consistent with the conclusions of Zak et al. (1994),

who suggested that a correlation between microbial biomass

and plant production among North American sites ranging

from deserts to temperate forests was evidence that carbon

Table 3

Microbial respiration rates and biomass pools in ‘control’ soil samples (i.e. only H2O was added) from three loblolly pine stands in North Carolina

Site 1 Site 2 Site 3

Forest floor

CO2-C respireda (mg C g soil21 week21) 7407 (1030)b 5083 (530)c 8486 (741)a

Biomass Cb (mg C g soil21) 9285 (1210)a 11 284 (1540)a 12 868 (793)a

Biomass Nb (mg N g soil21) 595 (57)b 679 (91)ab 732 (110)a

Mineral Soilc

CO2-C respireda (mg C g soil21week21) 210 (56)a 174 (17)a 257 (59)a

Biomass Cb (mg C g soil21) 633 (101)a 334 (16)b 396 (102)b

Biomass Nb (mg N g soil21) 94.3 (6.6)a 36.7 (7.0)c 56.9 (13.5)b

Data shown are means of four observations per site, with the standard errors in parentheses. These standard errors were calculated using only data from the

four control samples from each site. Values in a row with the same letter are not significantly different at P , 0:05 (Scheffe’s Tests, derived from ANOVAs that

used all 96 observations).a Respiration integrated over 7-day laboratory incubation at 22 8C.b Measured after 7-day laboratory incubation at 22 8C. Biomass was determined using a fumigation–extraction method. Chloroform-labile N and C were

divided by KEN ¼ 0:54 or KEC ¼ 0:45 to determine microbial biomass pools.c Intact cores from 0 to 7.5 cm depth.

Fig. 5. Microbial biomass C in intact cores of mineral soil (0–7.5 cm depth)

from three loblolly pine stands in central North Carolina, 7 days after

applying C, N or P solutions alone or in combination. Data are averaged

across two sampling dates and three sites. Microbial biomass C was

determined by chloroform fumigation–extraction, and dissolved organic C

values (determined by catalyzed combustion) were divided by a KEC

correction coefficient of 0.45 to convert to biomass values. Error bars are 1

standard error, calculated using only the 12 values for a given treatment.

These error bars include some variability due to differences among sites and

dates that is accounted for in the ANOVA used to produce the Scheffe

statistics. Bars with the same letter are not significantly different (P . 0:05;

Scheffe’s Test).

A.S. Allen, W.H. Schlesinger / Soil Biology & Biochemistry 36 (2004) 581–589 587

often limits soil microbes. The response of microbial

biomass to C addition may indicate a potential for microbial

biomass to respond to increased C inputs from plants grown

at elevated atmospheric CO2, as described by Zak et al.

(1993) and Dıaz et al. (1993).

We found that addition of NH4NO3 to mineral soil

significantly increased microbial biomass N (Fig. 6), and

that microbial biomass C and respiration tended to be higher

with NH4NO3 addition (Figs. 4 and 5). Tracer studies in

which 15NH4þ is added to forest soil typically find a large

proportion of 15N in microbial biomass or microbial

byproducts (Vitousek and Matson, 1984; Zak et al., 1990).

In light of numerous observations of microbial stimulation

by labile C addition (i.e. Anderson and Domsch, 1985),

these observations may be indicative of colimitation of

microbes by C and N. A second plausible explanation is that

several distinct populations exist within microbial biomass,

of which one is C-limited and another is N-limited (Cochran

et al., 1988). Our results are also consistent with those of

Hart and Stark (1997), who found that N additions increased

microbial biomass N in the 4–10 cm layer of mineral soil of

an old-growth, mixed conifer forest.

Our results differ from those of Gallardo and Schlesinger

(1994) and Scheu (1990), who found P limitation to

microbial biomass N in the mineral soil of temperate

Hardwood forests. Gallardo and Schlesinger (1994) suggest

that in late-successional forests on highly weathered, acid

soils, large quantities of P may be sequestered in Al or Fe

sesquioxides or in plants. In the present study, all three sites

had been severely disturbed 16 years prior to the time of our

measurements (Table 1), and P availability may have been

relatively high.

Addition of N to mineral soil samples in the absence of C

addition consistently decreased the C:N ratio in microbial

biomass while having little effect on respiration. This result

suggests that microbes took up N beyond their current

metabolic requirements (i.e. ‘luxury consumption’), or that

N addition caused a shift in dominance of the microbial

community from microbes with a high C:N ratio (i.e. fungi)

to microbes with a low C:N ratio (i.e. bacteria; Paul and

Clark, 1989).

5. Conclusions

We found that additions of C dramatically accelerated

respiration in both mineral soil and forest floor material, and

that C addition increased microbial biomass C in the mineral

soil. Additions of N increased respiration in forest floor

material and increased microbial biomass N in the mineral

soil. Addition of P caused a small increase in forest floor

respiration, but had no effect on microbial biomass. Our

results suggest that any additional inputs to soil of labile C

due to plant growth at elevated CO2 will result in increased

soil respiration and microbial biomass C. These results are

not consistent with the hypothesis that higher C:N ratios in

litter will decrease decomposition rates. However, our

results are consistent with the hypothesis that microbial

activity increases when the C supply to microbes is

increased. Our observations of increased respiration in

forest floor samples to which N was added suggest that

increases in atmospheric N deposition may accelerate

decomposition of forest floor material in loblolly pine

forests.

Acknowledgements

Many thanks go to Heavin Bortz for help in the field and

the laboratory. Jeffrey Andrews and Heather Hemric gave

important technical assistance. Judson Edeburn and Wendy

Weiher provided maps and land-use information. Donald

Burdick patiently provided statistical advice. James Clark,

Boyd Strain, Daniel Richter and Michael Lavine made

useful comments on an earlier draft of the manuscript.

Andrew S. Allen was supported by a NASA Earth System

Science Fellowship.

References

Aber, J., McDowell, W., Nadelhoffer, K., Magill, A., Berntson, G.,

Kamakea, M., McNulty, W., Rustad, L., Fernandez, I., 1998. Nitrogen

saturation in temperate forest ecosystems. BioScience 48, 921–934.

Allen, A.S., 1999. Effects of elevated atmospheric CO2 on soil nitrogen

availability. Dissertation. Duke University, Durham, NC, USA.

Anderson, T., Domsch, K., 1985. Determination of ecophysiological

maintenance carbon requirements of soil microorganisms in a dormant

state. Biology and Fertility of Soils 1, 81–89.

Fig. 6. Microbial biomass N in intact cores of mineral soil (0–7.5 cm depth)

from three loblolly pine stands in central North Carolina, 7 days after

applying C, N or P solutions alone or in combination. Data are averaged

across two sampling dates and three sites. Microbial biomass C was

determined by chloroform fumigation–extraction, and dissolved organic C

values (determined by catalyzed combustion) were divided by a KEN

correction coefficient of 0.54 to convert to biomass values. Error bars are 1

standard error, calculated using only the 12 values for a given treatment.

These error bars include some variability due to differences among sites that

is accounted for in the ANOVA used to produce the Scheffe statistics. Bars

with the same letter are not significantly different (P . 0:05; Scheffe’s

Test).

A.S. Allen, W.H. Schlesinger / Soil Biology & Biochemistry 36 (2004) 581–589588

Berg, B., 1986. Nutrient release from litter and humus in coniferous forest

soils—a mini review. Scandinavian Journal of Forest Research 1,

359–369.

Brookes, P.C., Landman, A., Pruden, G., Jenkinson, D.S., 1985. Chloro-

form fumigation and the release of soil nitrogen: a rapid direct

extraction method to measure microbial biomass nitrogen in soil. Soil

Biology & Biochemistry 17, 837–842.

Cochran, V.L., Horton, K.A., Cole, C.V., 1988. An estimation of

microbial death rate and limitations of nitrogen or carbon during

wheat straw decomposition. Soil Biology & Biochemistry 20,

293–298.

Curtis, P.S., Wang, X., 1998. A meta-analysis of elevated CO2 effects

on woody plant mass, form, and physiology. Oecologia 113,

299–313.

D’Elia, C.F., Steudler, P.A., Corwin, N., 1977. Determination of total

nitrogen in aqueous samples using persulfate oxidation. Limnology and

Oceanography 22, 760–764.

DeLucia, E.H., Hamilton, J.G., Naidu, S.L., Thomas, R.B., Andrews,

J.A., Finzi, A., Lavine, M., Matamala, R., Mohan, J.E., Hendrey,

G.R., Schlesinger, W.H., 1999. Net primary production of a forest

ecosystem under experimental CO2 enrichment. Science 284,

1177–1179.

Dıaz, S., Grime, J.P., Harris, J., McPherson, E., 1993. Evidence of a

feedback mechanism limiting plant response to elevated carbon

dioxide. Nature 364, 616–617.

Ellsworth, D.S., 1999. CO2 enrichment in a maturing pine forest: are CO2

exchange and water status in the canopy affected? Plant Cell and

Environment 22, 461–472.

Finzi, A.C., Allen, A.S., DeLucia, E.H., Ellsworth, D.S., Schlesinger,

W.H., 2001. Forest litter production, chemistry, and decomposition

following two years of free-air CO2 enrichment. Ecology 82,

470–484.

Gallardo, A., Schlesinger, W.H., 1990. Estimating microbial biomass

nitrogen using the fumigation–incubation and fumigation–extraction

methods in a warm-temperate forest soil. Soil Biology & Biochemistry

22, 927–932.

Gallardo, A., Schlesinger, W.H., 1994. Factors limiting microbial biomass

in the mineral soil and forest floor of a warm-temperate forest. Soil

Biology & Biochemistry 26, 1409–1415.

Hart, S.C., Stark, J.M., 1997. Nitrogen limitation of the microbial biomass

in an old-growth forest soil. Ecoscience 4, 91–98.

Hart, S.C., Nason, G.E., Myrold, D.D., Perry, D.A., 1994. Dynamics of

gross nitrogen transformations in an old-growth forest: the carbon

connection. Ecology 75, 880–891.

Horwath, W.R., Paul, E.A., 1994. Microbial biomass. In: Weaver, R.W.,

Angle, J.S., Bottomley, P.S. (Eds.), Methods of Soil Analysis, Part 2.

Microbiological and Biochemical Properties, Madison, WI, pp.

753–773.

Hungate, B.A., Holland, E.A., Jackson, R.B., Chapin, F.S. III, Mooney,

H.A., Field, C.B., 1997. The fate of carbon in grasslands under carbon

dioxide enrichment. Nature 388, 576–579.

Jenkinson, D.S., Powlson, D.S., 1976. The effects of biocidal treatments on

metabolism in soil—I. Fumigation with chloroform. Soil Biology &

Biochemistry 8, 167–177.

Jonasson, S., Vestergaard, P., Jensen, M., Michelsen, A., 1996a.

Effects of carbohydrate amendments on nutrient partitioning, plant

and microbial performance of a grassland–shrub ecosystem. Oikos

75, 220–226.

Jonasson, S., Michelsen, A., Schmidt, I.K., Nielsen, E.V., Callaghan, T.V.,

1996b. Microbial biomass C, N and P in two arctic soils and responses

to addition of NPK fertilizer and sugar: implications for plant nutrient

uptake. Oecologia 106, 507–515.

Lichter, J., Lavine, M., Mace, K.A., Richter, D.D., Schlesinger, W.H.,

2000. Throughfall chemistry in a loblolly pine plantation under elevated

atmospheric CO2 concentrations. Biogeochemistry 50, 73–93.

Magill, A.H., Aber, J.D., 1998. Long-term effects of experimental nitrogen

additions on foliar litter decay and humus formation in forest

ecosystems. Plant and Soil 203, 301–311.

Matamala, R., Schlesinger, W.H., 2000. Effects of elevated atmospheric

CO2 on fine root production and activity in an intact temperate forest

ecosystem. Global Change Biology 6, 967–979.

Melillo, J.M., Aber, J.D., Muratore, J.F., 1982. Nitrogen and lignin control

of Hardwood leaf litter decomposition dynamics. Ecology 63,

621–626.

Myrold, D.D., 1987. Relationship between microbial biomass nitrogen and

a nitrogen availability index. Soil Science Society of America Journal

51, 1047–1049.

NADP (National Atmospheric Deposition Program) 1999. NRSP-3

[Online]. Available by National Trends Network, http://nadp.sws.

uiuc.edu/.

Paul, E.A., Clark, F.E., 1989. Soil Microbiology and Biochemistry.

Academic Press, San Diego.

Scheu, S., 1990. Changes in microbial nutrient status during secondary

succession and its modification by earthworms. Oecologia 84,

351–358.

Schlesinger, W.H., 1997. Biogeochemistry: An Analysis of Global Change.

Academic Press, San Diego, CA, p. 588.

Sierra, J., Renault, P., 1995. Oxygen consumption by soil microorganisms

as affected by oxygen and carbon dioxide levels. Applied Soil Ecology

2, 175–184.

Tietema, A., 1998. Microbial carbon and nitrogen dynamics in coniferous

forest floor material collected along a European nitrogen deposition

gradient. Forest Ecology and Management 101, 29–36.

Vance, E.D., Brookes, P.C., Jenkinson, D.S., 1987. An extraction method

for measuring soil microbial biomass C. Soil Biology & Biochemistry

19, 703–707.

Vitousek, P.M., Matson, P.A., 1984. Mechanisms of nitrogen retention in

forest ecosystems: a field experiment. Science 225, 51–52.

Wardle, D.A., 1992. A comparative assessment of factors which influence

microbial biomass carbon and nitrogen levels in soil. Biological Review

67, 321–358.

Zak, D.R., Groffman, P.M., Pregitzer, K.S., Christensen, S., Tiedje, J.M.,

1990. The vernal dam: plant-microbe competition for nitrogen in

Northern Hardwood forests. Ecology 71, 651–656.

Zak, D.R., Pregitzer, K.S., Curtis, P.S., Teeri, J.A., Fogel, R., Randlett,

D.L., 1993. Elevated CO2 and feedback between carbon and nitrogen

cycles. Plant and Soil 151, 105–117.

Zak, D.R., Tilman, D., Parmenter, R.R., Rice, C.W., Fisher, R.M., Vose, J.,

Milchunas, D., Martin, C.W., 1994. Plant production and soil

microorganisms in late-successional ecosystems: a continental-scale

study. Ecology 75, 2333–2347.

A.S. Allen, W.H. Schlesinger / Soil Biology & Biochemistry 36 (2004) 581–589 589