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APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Nov. 2011, p. 7499–7507 Vol. 77, No. 21
Copyright © 2011, American Society for Microbiology. All Rights Reserved.
Microbial Dynamics during Aerobic Exposure of Corn Silage Stored
under Oxygen Barrier or Polyethylene Films
and Giorgio Borreani
Dipartimento di Valorizzazione e Protezione delle Risorse Agroforestali, Agricultural Microbiology and Food Technology Sector,
University of Torino, Via L. da Vinci 44, 10095 Grugliasco (Torino), Italy,
and Dipartimento di Agronomia, Selvicoltura e
Gestione del Territorio, University of Torino, Via L. da Vinci 44, 10095 Grugliasco (Torino), Italy
Received 8 April 2011/Accepted 29 July 2011
The aims of this study were to compare the effects of sealing forage corn with a new oxygen barrier ﬁlm with
those obtained by using a conventional polyethylene ﬁlm. This comparison was made during both ensilage and
subsequent exposure of silage to air and included chemical, microbiological, and molecular (DNA and RNA)
assessments. The forage was inoculated with a mixture of Lactobacillus buchneri,Lactobacillus plantarum, and
Enterococcus faecium and ensiled in polyethylene (PE) and oxygen barrier (OB) plastic bags. The oxygen
permeability of the PE and OB ﬁlms was 1,480 and 70 cm
per 24 h at 23°C, respectively. The silages were
sampled after 110 days of ensilage and after 2, 5, 7, 9, and 14 days of air exposure and analyzed for
fermentation characteristics, conventional microbial enumeration, and bacterial and fungal community ﬁn-
gerprinting via PCR-denaturing gradient gel electrophoresis (DGGE) and reverse transcription (RT)-PCR-
DGGE. The yeast counts in the PE and OB silages were 3.12 and 1.17 log
, respectively, with
corresponding aerobic stabilities of 65 and 152 h. Acetobacter pasteurianus was present at both the DNA and
RNA levels in the PE silage samples after 2 days of air exposure, whereas it was found only after 7 days in the
OB silages. RT-PCR-DGGE revealed the activity of Aspergillus fumigatus in the PE samples from the day 7 of
air exposure, whereas it appeared only after 14 days in the OB silages. It has been shown that the use of an
oxygen barrier ﬁlm can ensure a longer shelf life of silage after aerobic exposure.
Forage ensiling is based on the natural fermentation of wa-
ter-soluble plant carbohydrates by lactic acid bacteria (LAB)
under anaerobic conditions (24). The most important single
factor that can inﬂuence the preservation efﬁciency of forage
ensiling is the degree of anaerobiosis reached in the completed
silo (36). Anaerobic conditions are not always achieved in silos
on individual farms, especially in the outer layer of a silo,
because of the difﬁculty of sealing it efﬁciently (6). The aerobic
deterioration of silages is a signiﬁcant problem for farm prof-
itability and feed quality throughout the world. All silages
exposed to air deteriorate as a result of aerobic microbial
activity during feed-out (8, 31, 35). These losses can reach 70%
of the stored dry matter in the top layer and near the sidewalls
of the bunkers and are related to the depletion of the digestible
carbohydrate and organic acid fractions (5). Spoilage of silage
due to exposure to air is undesirable, due to the resulting
decrease in nutritive value and to the risk of negative effects on
animal performance (18), which are also connected to the
proliferation of potentially pathogenic or otherwise undesir-
able microorganisms (20) and mycotoxin synthesis (28).
Polyethylene (PE) ﬁlms have been used for many years to
seal bunker silos and drive-over piles because of their suitable
mechanical characteristics and low costs. The high O
ability of PE ﬁlms can contribute to the low quality of silage in
the top layer of horizontal silos (6). A new silage sealing plastic
ﬁlm, which uses a new plastic formulation with an 18-fold-
lower oxygen permeability than the PE ﬁlm usually used on
farms, has recently been developed (7). Polymers different
from PE, such as polyamides (PA) and ethylene-vinyl alcohol
(EVOH), help create an excellent barrier against oxygen, com-
bined with good mechanical characteristics (puncture resis-
tance), and are suitable for blown coextrusion with PE to
produce 45- to 200-m-thick plastic ﬁlms.
Monitoring microbiota during the ensiling process has be-
come more reliable and accurate, thanks to recently developed
culture-independent methods (25, 29). In recent years, DNA-
based community ﬁngerprinting techniques, such as denatur-
ing gradient gel electrophoresis (DGGE) and terminal restric-
tion fragment length polymorphism (T-RFLP), have been
applied to investigate the microbial community composition of
silage (9, 22). Advanced molecular biological techniques have
been used to further our understanding of the structure of
complex microbial community dynamics (25). However, to the
best of our knowledge, no study has used a community ﬁnger-
printing approach to investigate the population dynamics of
corn silage during aerobic deterioration.
In this study, a culture-independent technique, PCR-
DGGE, was used to study microbial dynamics, whereas reverse
transcription (RT)-PCR-DGGE allowed us to investigate the
metabolically active populations. These techniques were per-
formed together with conventional microbiological enumera-
tion in order to investigate the effects of a new oxygen barrier
ﬁlm on the microbial community of ensiled and aerobically
deteriorated corn silages.
MATERIALS AND METHODS
Crop and ensiling. The trial was carried out at the experimental farm of the
University of Turin on the western Po plain in northern Italy (44°50⬘N, 7°40⬘E;
* Corresponding author. Mailing address: Dipartimento di Agrono-
mia, Selvicoltura e Gestione del Territorio, University of Torino, Via
L. da Vinci 44, 10095 Grugliasco (Torino), Italy. Phone: (39) 011
6708783. Fax: (39) 011 6708798. E-mail: firstname.lastname@example.org.
Published ahead of print on 5 August 2011.
altitude, 232 m above sea level) in 2008 on corn (Zea mays L.) harvested as a
whole-corn crop, at a 50% milk line stage, and at 333 g dry matter (DM) kg
fresh forage. The forage was chopped with a precision forage harvester to a
10-mm theoretical length and inoculated with a mixture of Lactobacillus buchneri
(strain ATCC PTA2494), Lactobacillus plantarum (strains ATCC 53187 and
55942), and Enterococcus faecium (strain ATCC 55593) (inoculum 11C33; Pio-
neer Hi-Bred International, Des Moines, IA) to yield 1 ⫻10
CFU per gram of fresh forage, respectively. Standard black-on-white poly-
ethylene ﬁlm, 120 m thick (PE), and 120-m-thick Silostop (Bruno Rimini Ltd.,
London, United Kingdom), black-on-white coextruded polyethylene-polyamide
ﬁlm with an enhanced oxygen barrier (OB), were used to produce the silage bags
for this experiment. Bags were heat sealed at the closed end and were equipped
with a one-way valve for CO
release. Each bag was inserted into a portion of a
PVC (polyvinyl chloride) tube (internal dimensions, 300-mm diameter and
300-mm height; 21-liter volume) so that just the top and the bottom of the bag
had access to air. All bags were then ﬁlled with about 12 kg of fresh forage, which
was compacted manually, and secured with plastic ties. Four replicates were
prepared for each treatment. The density of the silage was 576 kg fresh matter
and 192 kg DM m
. The oxygen permeability of PE and OB,
determined by ASTM standard method D 3985-81 (4), was 1,480 and 70 cm
per 24 h at 100 KPa at 23°C and 0% relative humidity, respectively. The silos
were stored at ambient temperature (18 to 22°C) indoors and opened after 110
days. The ﬁnal weights were recorded at silo opening, and the silage was mixed
thoroughly and subsequently sampled. The DM concentration (three replicates)
and fermentation end products (two replicates) were determined for each sam-
ple. Microbiological counts (two replicates) and culture-independent techniques
(two replicates) were also carried out. The silages were subjected to an aerobic
stability test. Aerobic stability was determined by monitoring the temperature
increases due to the microbial activity of the samples exposed to air. About three
kilograms from each silo was allowed to aerobically deteriorate at room temper-
ature (22 ⫾1.6°C) in 17-liter polystyrene boxes (290-mm diameter and 260-mm
height) for 14 days. A single layer of aluminum cooking foil was placed over each
box to prevent drying and dust contamination but also allowed air penetration.
The temperature of the room and of the silage was measured each hour by a data
logger. Aerobic stability was deﬁned as the number of hours the silage remained
stable before rising more than 2°C above room temperature (27). The silage was
sampled after 0, 2, 5, 7, 9 and 14 days of aerobic exposure to quantify the
microbial and chemical changes of the silage during exposure to air.
Sample preparation and analyses. Each of the pre-ensiled samples of each
herbage and the samples of silage taken from each bag of silage were split into
three subsamples. One subsample was oven-dried at 65°C to constant weight to
determine the DM content and air equilibrated, weighed, and ground in a
Cyclotec mill (Tecator, Herndon, VA) to pass through a 1-mm screen. The dried
samples were analyzed for total nitrogen (TN) by combustion (30), according to
the Dumas method, using a Micro-N nitrogen analyzer (Elementar, Hanau,
Germany), and for ash by complete combustion in a mufﬂe furnace at 550°C for
3 h. A portion of the second subsample was extracted using a stomacher blender
(Seward Ltd., London United Kingdom) for 4 min in distilled water at a ratio of
water to sample material (fresh weight) of 9:1, and another portion was extracted
(0.05 mol liter
) at a ratio of acid to sample material (fresh weight)
of 5:1. The nitrate (NO
) contents were determined in the water extract, through
semiquantitative analysis, using Merckoquant test strips (7). The ammonia ni-
-N) content, determined using a speciﬁc electrode, was quantiﬁed in
the water extract. The lactic and monocarboxylic acids (acetic, propionic, and
butyric acids) were determined by high-performance liquid chromatography
(HPLC) in the acid extract (10). Ethanol, for which the HPLC was coupled to a
refractive index detector, was also measured using an Aminex HPX-87H column
(Bio-Rad Laboratories, Richmond, CA). The analyses were performed isocrati-
cally under the following conditions: mobile phase, 0.0025 mol liter
ﬂow rate, 0.5 ml min
; column temperature, 37°C; injection volume, 100 l.
Duplicate analyses were performed for all the determined parameters. The
duplicates were averaged, and the four means (one for each silo) were consid-
ered four observations in the statistical analysis. The water activity (a
silage was measured at 25°C using an AquaLab series 3TE instrument (Decagon
Devices Inc., Pullman, WA) on a fresh sample at silo opening. The weight losses
due to fermentation were calculated as the difference between the weight of the
plant material placed in each silo at ensiling and the weight of the silage at the
end of conservation.
A third subsample was used for the microbiological analyses. For the microbial
counts, 30 g of sample were transferred into sterile homogenization bags, sus-
pended at 1:10 (wt/vol) in peptone salt solution (PPS;1gofbacteriological
peptone and9gofsodium chloride per liter), and homogenized for 4 min in a
stomacher blender (Seward Ltd., London, United Kingdom). Serial dilutions
were prepared, and the following counts were carried out: (i) aerobic spores after
pasteurization at 80°C for 10 min followed by double-layer pour plating with
24.0 g liter
nutrient agar (NUA; Oxoid, Milan, Italy) and incubation at 30°C
for 3 days; (ii) mold and yeast on 40.0 g liter
of yeast extract glucose chlor-
amphenicol agar (YGC agar; Difco, West Molesey, Surrey, United Kingdom)
after incubation at 25°C for 3 and 5 days for yeast and mold, respectively. The
mean count of the duplicate subsamples was recorded for the microbial counts
on plates that yielded 10 to 100 CFU per petri dish.
Microbial community ﬁngerprinting by PCR-DGGE and RT-PCR-DGGE. (i)
Sampling and nucleic acid extraction. Two milliliters of the supernatant of the
above-described 1:10 diluted sample suspension were collected for each sampling
point and centrifuged at 13,400 ⫻gfor 5 min to pellet the cells. After the
supernatant had been discarded, the pellet was subjected to DNA and RNA
extraction using DNeasy and RNeasy plant minikits (Qiagen, Milan, Italy),
respectively, according to the manufacturer’s instructions. The presence of re-
sidual DNA in the RNA samples was checked by PCR (12).
(ii) PCR and RT-PCR. The dominant bacterial microbiota was investigated, at
both the DNA and RNA level, by PCR-DGGE and RT-PCR-DGGE. The
primers 338fGC and 518r were used to detect and amplify the bacterial variable
region of 16S rRNA gene (1). In order to investigate the dominant fungal
microbiota, the D1-D2 loop of the 26S rRNA gene was ampliﬁed by PCR using
the primers NL1GC and LS2 (11).
(iii) DGGE analysis. The Dcode universal mutation detection system (Bio-
Rad) was used to perform DGGE analysis. The amplicons obtained from the
PCR and RT-PCR were applied to an 8% (wt/vol) polyacrylamide gel (acryl-
amide-bisacrylamide, 37.5:1) with a 30%-to-60% denaturant gradient (13). Some
selected DGGE bands were excised from the gels and incubated overnight at 4°C
in 50 l of sterile water. The eluted DNA was reampliﬁed and analyzed in
DGGE (1). The amplicons that gave a single band comigrating with the control
were then ampliﬁed with a 338f primer and NL1 primer, respectively, for bac-
terial and fungal microbiota without a GC clamp and puriﬁed with Perfectprep
gel clean-up (Eppendorf, Milan, Italy) for sequencing.
(iv) Sequence analysis. The PCR-DGGE and RT-PCR-DGGE bands were
sent for sequencing to Euroﬁns MWG Operon (Ebersberg, Germany), and the
gene sequences obtained were aligned with those in GenBank using the BLAST
program (2) to establish the closest known relatives of the amplicons run in
Statistical analysis. All microbial counts and hours of aerobic stability were
transformed to obtain log-normal-distributed data. The fermentative char-
acteristics, microbial counts, pH, nitrate contents, dry weight losses, and hours of
aerobic stability were subjected to a one-way analysis of variance (Statistical
Package for Social Science, version 16; SPSS Inc., Chicago, IL) to evaluate the
statistical signiﬁcance of the differences between the two treatments. Between-
treatment comparisons were made using an unpaired Student’s ttest, and dif-
ferences were considered signiﬁcant at P⬍0.05.
DGGE proﬁles were normalized and subjected to cluster analysis using
BioNumerics software (Applied Maths, Kortrijk, Belgium). The Pearson product
moment correlation coefﬁcient was used to calculate the similarities in DGGE
patterns, and dendrograms were obtained via the unweighted pair group method
with arithmetic averages.
Fermentative quality and microbial counts of the silages.
The results of the chemical and microbial determination of the
corn forage prior to ensiling are shown in Table 1. The values
are typical of those of corn harvested at a 50% milk line. The
fermentation quality and microbial composition of the silages,
after 110 days of conservation, are shown in Table 2. All the
silages were well fermented. The main fermentation acids
found were lactic and acetic acids, whereas butyric acid was
below the detection limit (less than 0.1 g kg
DM) in all the
silages. The silages sealed with the PE ﬁlm led to silages with
higher pH (P⬍0.002), and lower concentrations of lactic acid
(P⬍0.033) in comparison to the OB silages. The nitrate levels
in the corn crop were lower in the silages than in the corre-
sponding herbage. The yeast counts were lower below the OB
ﬁlm, whereas the mold count was below 2 log
in both treatments. The a
of the silages at opening had a
7500 DOLCI ET AL. APPL.ENVIRON.MICROBIOL.
mean value of 0.99, and there was no difference between the
two treatments. The weight losses were lower in the OB silages
than in the PE silages. The aerobic stabilities of the silages ex-
posed to air were 65 and 152 h in the PE and in OB silages,
Silage quality during the air exposure test. The changes in
temperature, pH, lactic acid, yeast and mold counts, and num-
bers of aerobic spores in the silages for 14 days of aerobic
exposure are reported in Fig. 1. The temperature at silo open-
ing was about 22°C for both treatments. After 65 h of aerobic
exposure, the temperature of the PE silages started to rise,
whereas the temperature in the OB silages did not increase
over the ﬁrst 6 days of exposure to air. The PE silages showed
temperatures above 35°C after 4.7 days (113 h) and reached
the highest temperature of 42.2°C after 9.5 days (229 h). Under
the OB ﬁlm, temperatures above 35°C were reached only after
11.4 days (274 h) of air exposure. Simultaneously with the
variation in silage temperatures, a pH increase was observed in
the two treatments. The pH was always lower in the OB than
in the PE silages. The lactic acid concentration started to
decrease after 2 days in the PE silage and after 7 days in the
OB silage. The yeast count increased from the second day of
air exposure in both treatments and reached 6 log
silage after 5 days of air exposure. The mold counts remained
almost constant till day 5 of air exposure and started to in-
crease at day 7, with higher values in the PE silage than in the
OB silage. They reached similar values after 14 days of air
exposure. The aerobic spore count increased with air exposure
time in the PE silages, reaching a maximum value of 9.3 log
after 14 days of air exposure. The aerobic spore
count in the OB silages remained almost constant till day 9 of
air exposure and reached a value of 7.8 log
days of air exposure.
Bacterial community ﬁngerprinting of the silages and
aerobically exposed silages. The bacterial microbiota dynamics
was well described through the PCR-DGGE and RT-PCR-
DGGE proﬁles. The DNA and RNA gels are shown in Fig. 2,
and the band identiﬁcation results are reported in Table 3. The
inoculated starter, L. buchneri, was present for 14 days, at the
DNA level, in the OB silage samples (Fig. 2c, band a), whereas
it was found for 5 days in the PE silages (Fig. 2a, band a). L.
plantarum was detected in the samples ensiled in OB only
between days 5 and 14 (Fig. 2c, band g). Faint bands were
found at the RNA level for these two species. L. buchneri was
not detected beyond 7 days (Fig. 2d, band a), and L. plantarum
was detected in the OB samples from days 5 to 9 (Fig. 2d, band
g). Acetobacter pasteurianus was clearly present at both DNA
and RNA levels in the PE silage samples, except for the days
immediately subsequent to air exposure (Fig. 2a and b, band
b). A. pasteurianus was found in OB silages only after 7, 9, or
14 days of aerobic exposure (Fig. 2c and d, band b). Faint
bands corresponding to Bacillus subtilis were detected at the
DNA level in the PE silage samples from day 5 to day 14 (Fig.
2a, band c), whereas B. subtilis was recovered in the DNA
extracted from the OB samples and at the RNA level only at
day 14 (Fig. 2b, c, and d, band c). Lactobacillus amylovorus was
found at the RNA level in PE silage upon opening (Fig. 2b,
band d), together with a band corresponding to an uncultured
bacterium (Fig. 2b, band e). Finally, a more persistent band,
again identiﬁed by sequencing as uncultured bacterium, was
observed in the same samples, from day 5 to day 14.
Fungal community ﬁngerprinting of the silages and aerobi-
cally exposed silages. The fungal population was detected at
the DNA level, in both the PE and OB silages, with a band
present from opening to day 7 (Fig. 3a and c, band i, and Table
4). After sequencing, the band was determined to be Kazach-
stania exigua.Aspergillus fumigatus was found after 14 days of
air exposure (Fig. 3a and c, band h). DNA bands correspond-
ing to Pichia kudriavzevii were revealed in PE silages at day 7
and day 9 (Fig. 3a, band l). RT-PCR-DGGE revealed activity
of A. fumigatus, in particular in the PE silages, where it was
detected from day 7 to day 14 of air exposure (Fig. 3b, band h).
Unlike the DNA analysis, RNA identiﬁed a new band, which
was sequenced as Aureobasidium pullulans. This species was
present for 14 days of aerobic exposure in OB silage (Fig. 3d,
TABLE 1. Chemical and microbial composition of the corn forage
prior to ensiling
DM (g kg
) .................................................................................. 333
TN (g kg
DM) ........................................................................... 12.6
Starch (g kg
NDF (g kg
DM) ........................................................................ 443
ADF (g kg
DM) ........................................................................ 248
Ash (g kg
DM) .......................................................................... 39.0
Nitrate (mg kg
herbage) ................................................. 6.13
Aerobic spores (log
herbage) .................................. 3.57
, water activity; ADF, acid detergent ﬁber; DM, dry matter; NDF, neutral
detergent ﬁber; TN, total nitrogen.
TABLE 2. Fermentation quality and microbial composition at
unloading of silages sealed with oxygen barrier (OB) and
standard polyethylene (PE) ﬁlms after 110
days of conservation
Value for: SE Pvalue
pH 3.78 3.73 0.011 0.002
DM (g kg
)297 310 8.22 0.500
Lactic acid (g kg
DM) 45.3 53.2 2.08 0.033
Acetic acid (g kg
DM) 27.6 22.7 1.23 0.019
Butyric acid (g kg
DM) ⬍0.10 ⬍0.10
Propionic acid (g kg
DM) 0.45 0.76 0.164 0.402
1,2-Propanediol (g kg
DM) 10.5 10.4 0.706 0.981
Ethanol (g kg
DM) 12.3 11.2 0.651 0.443
Lactic-to-acetic acid ratio 1.64 2.34 0.165 0.004
Nitrate (mg kg
silage) 837 1026 156 0.603
-N (g kg
TN) 46.8 45.8 0.112 0.746
Ash (g kg
DM) 40.6 40.2 0.031 0.656
0.99 0.99 0.001 0.947
silage) 3.12 1.17 0.443 ⬍0.001
silage) 1.74 1.41 0.118 0.189
Aerobic spores (log
2.65 2.97 0.095 0.095
Weight loss (g kg
DM) 37.5 30.6 0.178 0.035
Aerobic stability (h) 65 152 19.9 0.001
, water activity; C, control treatment; DM, dry matter; LAB, lactic acid
-N, ammonia nitrogen; TN, total nitrogen.
VOL. 77, 2011 AEROBIC SPOILAGE OF CORN SILAGE UNDER DIFFERENT FILMS 7501
band m), whereas it was found for only 5 days of air exposure
in the PE silages (Fig. 3b, band m). Furthermore, a band that
could not be matched to any fungal species was present in OB
silages from day 2 to 9 (Fig. 3d, band n). Finally, at the DNA
level, bands run in the middle of the lanes were observed and
excised, but when they were reampliﬁed, unclear proﬁles were
obtained, and thus, they were interpreted as heteroduplexes.
No important differences in ﬁngerprints were observed be-
tween DNA and RNA among the replicates for each treat-
The cluster analysis highlighted the inﬂuence of the PE and
OB ﬁlms on the bacterial DGGE proﬁles (Fig. 4). A clustering
related to DNA and RNA analysis within each treatment can
be observed. The DGGE proﬁles of the mycobiota clustered in
two main groups, according to the nucleic acid analyzed (Fig.
5). Clusters correlated to both the sealing treatment and tem-
poral dynamics were noted at the RNA level.
An anaerobic environment is the most important individual
factor that can inﬂuence silage conservation (36). Most of the
FIG. 1. Dynamics of silage temperature, pH, lactic acid, yeast count, mold count, and aerobe spore count during air exposure of silages. PE,
polyethylene ﬁlm; OB, oxygen barrier ﬁlm.
7502 DOLCI ET AL. APPL.ENVIRON.MICROBIOL.
silages on individual farms are exposed to air during conser-
vation, due to the permeability of plastic to air and difﬁculties
in sealing the outer layer of silage properly, or during the
feed-out phase, due to an inadequate amount of silage being
removed and to a poor management of the exposed silo surface
(3). These observations highlight risks in terms of: spoilage
with losses in nutritional value (33), multiplication of poten-
tially pathogenic microorganisms, and production of mycotox-
ins (16). Since aerobic microbial populations increase during
aerobic deterioration in an exponential manner, the silages
from the spoiled top corner and from the molded spots have
the potential for contaminating feed-out silage to a great ex-
tent, even when it is included in very small amounts. To ad-
dress the issue of aerobic stability, inoculants containing L.
buchneri have been used over the last decade with the primary
purpose of increasing the amount of acetic acid and, as a
consequence, of decreasing yeast counts in silages (17). Plastic
oxygen barrier ﬁlms are also now available to cover silages and
to improve the anaerobic environment during conservation (6).
In our study, the fermentative proﬁles of silages stored un-
der both OB and PE ﬁlms were typical of fermentation driven
by L. buchneri, with a relatively high content of acetic acid, low
FIG. 2. DGGE proﬁles of bacterial DNA (a and c) and RNA (b and d) extracted from silage samples exposed to air for 0, 2, 5, 7, 9, and 14
days and ensiled in polyethylene plastic bags (PE) (a and b) or oxygen barrier bags (OB) (c and d). Letters indicate bands that were subjected to
sequencing as described in Materials and Methods, and the results are reported in Table 3. Lanes M, markers.
VOL. 77, 2011 AEROBIC SPOILAGE OF CORN SILAGE UNDER DIFFERENT FILMS 7503
lactic-to-acetic acid ratio (⬍3), and the presence of more than
DM of 1,2-propanediol. These values are in agree-
ment with those reported by Kleinschmit and Kung (17), who
summarized the effects of L. buchneri on silage quality in 43
experiments. Furthermore, the low permeability to oxygen of
the OB ﬁlm helped create a more anaerobic environment, and
this was reﬂected in a silage with a higher lactic acid content,
a lower pH and acetic acid content, and lower weight losses.
The better anaerobic environment under the OB ﬁlm also
contributed to reducing yeast counts to below 2.0 log
of silage. The reduction in yeast counts was reﬂected in an
TABLE 3. Sequence information for fragments detected on DGGE
gels obtained by analyzing the bacterial population through direct
DNA and RNA analysis of silage samples
Band Closest sequence relative Identity
aLactobacillus buchneri 99 HM162413
bAcetobacter pasteurianus 98 AP011156
cBacillus subtilis 100 HQ009797
dLactobacillus amylovorus 100 EF439704
e Uncultured bacterium 98 GU343612
f Uncultured bacterium 100 GQ233026
gLactobacillus plantarum 100 HQ117897
FIG. 3. DGGE proﬁles of fungal DNA (a and c) and RNA (b and d) extracted from silage samples exposed to air for 0, 2, 5, 7, 9, and 14 days
and ensiled in polyethylene plastic bags (PE) (a and b) or oxygen barrier bags (OB) (c and d). Letters indicate bands that were subjected to
sequencing as described in Materials and Methods, and the results are reported in Table 4. Lanes M, markers.
7504 DOLCI ET AL. APPL.ENVIRON.MICROBIOL.
increase in the aerobic stability of the OB silages, when ex-
posed to air. It is well known that lactate-assimilating yeasts
(Saccharomyces,Candida, and Pichia spp.) are generally the
main initiators of the aerobic spoilage of silages (24), and
under aerobic conditions, they utilize lactic acid, thus causing
an increase in silage temperature and pH. In our study, the
dominant yeast species after exposure to air, as observed from
the DGGE proﬁles of fungal DNA and RNA, was Kazachsta-
nia exigua, in both the PE and OB silages. Yeasts of the genus
of Kazachstania were previously observed in aerobically dete-
riorating corn silages (21). Furthermore, Pichia kudriavzevii
was observed in PE silages after 7 days of air exposure. Yeasts
of the genus Pichia are usually reported to be the initial cause
of aerobic deterioration of different silage crops (24). P.
kudriavzevii has recently been found in Italian ryegrass silage
treated with L. buchneri (19). From the DGGE proﬁles of
bacterial RNA at silage opening and during 14 days of air
exposure, apart from the presence of LAB, A. pasteurianus was
also seen to be present from the second day of air exposure in
the PE silages, while it appeared at day 7 in the OB silages.
This could partially explain the more rapid degradation that
occurred in the PE silage after exposure to air. Spoelstra et al.
(32) found that Acetobacter spp. could be involved in the aer-
obic spoilage of corn silage, by oxidizing ethanol to acetate or
by oxidizing lactate and acetate to carbon dioxide and water.
Furthermore, the selective inhibition of yeasts, due to the ad-
dition of acetic or propionic acid, could also increase the pro-
liferation of acetic acid bacteria in silage (15). Here, the use of
L. buchneri as a silage inoculant provoked a heterolactic fer-
mentation with an increase in the acetic acid concentration.
This could have indirectly stimulated the activity of A. pasteu-
rianus. The presence of A. pasteurianus in silages was recently
reported by Nishino et al. (23), who identiﬁed two strains of A.
pasteurianus in whole-crop corn silage which contained signif-
icant amounts of acetic acid and which had been stored for 18
When the yeasts and Acetobacter had consumed most of the
lactic acid (Fig. 1) and acetic acid (data not shown), the pH
level increased and the growth of other aerobic bacteria and
ﬁlamentous fungi became possible, which caused further spoil-
age (36). This secondary aerobic spoilage microbiota, which
principally consist of mold and bacilli, not only decreases the
nutritive value of the silage but also presents a risk to animal
health and the safety of milk (34). In this study, the aerobic
TABLE 4. Sequence information for fragments detected on DGGE
gels obtained by analyzing the fungal population through direct
DNA and RNA analysis of silage samples
Band Closest sequence relative Identity
hAspergillus fumigatus 99 HM807348
iKazachstania exigua 100 FJ468461
lPichia kudriavzevii 99 GQ894726
mAureobasidium pullulans 98 GQ281758
n Synthetic construct, ankyrin
repeat protein E2_17
gene, partial CDS
CDS, coding sequence.
FIG. 4. Dendrograms obtained from cluster analysis of DGGE proﬁles of the bacterial microﬂora detected on PE- and OB-treated silage
samples, at both the DNA and RNA levels, during aerobic exposure.
VOL. 77, 2011 AEROBIC SPOILAGE OF CORN SILAGE UNDER DIFFERENT FILMS 7505
spore counts increased above 8 log
from day 3 of
air exposure and beyond in the PE silage, whereas they tended
to increase in the OB silage only after 9 days of air exposure.
Bacillus sp. counts of up to 9 log
silage have been
detected in deteriorating silage and from the face layer of
opened bunker silages (24). The DGGE proﬁles showed that
B. subtilis was present in both the PE and OB silages. The
presence of A. fumigatus was observed after 7 days of aerobic
exposure in the PE silages and after 14 days in the OB silages,
when mold counts exceeded 6 log
silage. A. fumiga-
tus is a well-known human and animal pathogen that causes
aspergillosis, and it can produce gliotoxin, a toxic compound
that has potent immunosuppressive, genotoxic, cytotoxic, and
apoptotic effects (14, 26).
Overall, the cluster analysis highlighted the inﬂuence of PE
and OB ﬁlms on the metabolic activity of microbiota through-
out aerobic exposure. At the RNA level, clusters correspond-
ing to the sealing treatment were detected for both the bacte-
rial and fungal populations.
In this study, it has been shown that the use of oxygen barrier
plastic ﬁlms for ensiling can ensure a longer shelf life of silage,
protecting it from spoilage. Moreover, an important feature of
OB use is the delay in growth of pathogenic molds, which are
able to produce potent mycotoxins that are harmful to animals
This work was supported by the Regione Piemonte, Assessorato
Qualita`, Ambiente e Agricoltura years 2005–2008 Project: “Inﬂuenza
della zona di produzione e del tipo di gestione aziendale sulla qualita`
del Grana Padano D.O.P. piemontese.”
All the authors contributed equally to the work described in this
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