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NUTRITION, FEEDING, AND CALVES 
Computerized Monitoring of Gas Production to Measure 
Forage Digestion In Vitro 
A. N. PELL and P. SCHOFIELD 
Department of Animal Science 
Cornell University 
Ithaca, NY 14853 
ABSTRACT 
The techniques reported in this paper 
were developed to facilitate the study of 
the kinetics of forage digestion in vitro 
by measuring gas production. Fiber dis- 
appearance as a measure of the reaction 
rate has been replaced by the use of 
computerized pressure sensors to moni- 
tor the gaseous products ( C 0 2 , CH4) of 
microbial metabolism. The recording 
system described requires a computer, 
pressure sensors, an interface card, and 
appropriate software to monitor gas 
production continuously. Several varia- 
bles, including sample size, inoculum 
size, vessel size, and type of pressure 
sensor, have been investigated to deter- 
mine ranges within which gas production 
can be measured accurately, and the 
reproducibility of the results has been 
established. Because this technique uses 
small (100-mg) samples, a modified 
NDF method has been introduced that 
allows determination of extent of diges- 
tion at the end of an incubation in which 
gas production has been monitored. A 
strong linear relationship existed be- 
tween NDF disappearance and gas 
production. 
(Key words: in vitro, gas production, 
digestion rate) 
Abbreviation key: A/D = analog to digital, A: 
P = acetate to propionate ratio. 
INTRODUCTION 
Milk yield and growth of ruminants are 
limited largely by forage quality (20), and the 
importance of forage analysis to digestibility 
Received September 8, 1992. 
Accepted November 16, 1992. 
has long been recognized. The in vitro fermen- 
tation method of analysis (5 , 18) is widely 
used but has several disadvantages. 1) The 
fiber analysis destroys the sample, and a differ- 
ent sample is required for each time point. 
Studies of kinetics are, therefore, time- 
consuming, tedious, labor-intensive, and sub- 
ject to poor repeatability. 2) Early stages of 
digestion are difficult to study because the 
corresponding weight losses are small. 3) The 
role of soluble forage components cannot be 
determined. 4) Relatively large forage samples 
are needed, and the method is, therefore, diffi- 
cult to apply to materials available in limited 
quantities, such as tissue culture samples or 
isolated cell-wall fractions. 
As an alternative to measuring fiber disap- 
pearance gravimetrically, several researchers 
(4, 8, 10, 11, 12, 15, 16, 17, 19) haveusedgas 
production as a measure of carbon metabolism. 
Taya et al. (16) showed that gas production 
was related linearly to cellulose digestion in 
pure cultures of Ruminococcus albus, and 
Menke and Steingass (12) related gas produc- 
tion to feedstuff digestibility and energy value. 
Gas production has been measured manometri- 
cally (8, 10) or volumetrically (4, 12, 15) with 
manual recording of the results. 
An alternative way to measure gas produc- 
tion is to record pressure increases in a closed 
system. An automated system in which gas 
production was recorded on a strip chart was 
described by Wilkins in 1974 (23). In a recent 
preliminary report, Theodorou et al. (17) used 
an LED (light emitting diode) read-out pres- 
sure sensor that was inserted manually into 
each tube when measurements were taken. 
This paper describes a computerized gas 
production method. Individual pressure sensors 
that remain in place throughout the analysis 
were used to transmit data to an IBM- 
compatible computer via an analog to digital 
(A/D) card. As many data points as desired 
could be transmitted in an automated proce- 
1993 J Dairy Sci 76:1063-1073 1063 
1064 PELL AND SCHOFIELD 
A C 
INTERFACE 
CONTROLLER CONNECTOR 
PI- 
~. 
TENPERATURE 
SENSOR 
I A 4 - n H E A T E R PRESSLRE 
- 
I STIRRER 
- AC 
I 
A/D CARD 
COMPUTER 
BLOCK D I A G R A M 
Figure 1 . Block diagram showing connections among various components of the gas pressure measuring system. Only 
1 sensor is shown; 8 or 15 would normally be connected. The AC (alternating current) indicates the power source. The 
A/D card is the analog to digital card. 
dure with this system. The data can be readily 
transformed into a spreadsheet format for fur- 
ther analysis. A micro-NDF method, the rela- 
tionship between NDF digestion and gas 
production, and possible applications are dis- 
cussed. 
MATERIALS AND METHODS 
Equipment 
The system components (Figure 1) include 
an incubator with a multiplace stirrer, pressure 
sensors attached to the incubation flasks, an 
A/D converter card, and a computer with soft- 
ware. 
Incubator. Because of the need for multiple 
input-output lines, an incubator was con- 
structed from plywood and Styrofoam insula- 
tion. The heating element was a 60- or 
90-W light bulb controlled by an external 
homemade switching device that used a diode 
as a temperature sensor. Temperature stratifi- 
cation was minimized by two fans operating in 
the unit; one fan ran continuously, and the 
other fan was switched. The operating temper- 
ature could be adjusted via a trimmer poten- 
tiometer on the switching device and was set 
to 39°C. This temperature was maintained at 
39 f S"C, which is comparable with many 
commercially available incubators. Two AC 
(alternating current) outlet sockets were 
mounted on the incubator's interior back wall; 
one outlet was continuously live, and the other 
outlet was connected to the switching device 
and was live only when the heater lamp was 
on. 
Stirrer. Attempts to use a commercially 
available, electronic, multiplace, remotely 
powered stirrer inside the incubator were not 
successful. If operated continuously, the stirrer 
generated more heat, even in the intermittent 
stirring mode, than the incubator box could 
dissipate at the chosen operating temperature. 
Intermittent operation of the unit was frus- 
trated by the default reset design. Instead, a 
simple homemade device was constructed 
from timing motors (48 rpm; Number 2480; 
Journal of Dairy Science Vol. 76. No. 4. 1993 
COMPUTERIZED MONITORING OF GAS PRODUCTION 1065 
American Science & Surplus, Evanston, IL) 
and ceramic magnets (Number 43 12; American 
Science & Surplus). Each motor was con- 
figured to stir five sample bottles in a pen- 
tagonal array so that three motors would suf- 
fice for a 15-place stirrer. The motors were 
mounted in an open-frame assembly and were 
connected to the switched power supply. Stir- 
ring was thus intermittent and gentle and 
generated minimal heat. 
Pressure Sensors. Two types of sensors are 
available. Differential sensors provide a low 
full-scale voltage output (typically 50 to 100 
mV) and require dual output signal leads. A 
single-ended sensor has a built-in amplifier 
that raises the maximum signal level to about 5 
V; this type of sensor uses a single signal lead 
and is more expensive than its differential 
counterpart. We have used sensors of both 
types and prefer the single-ended type because 
it allows twice as many sensors to be inter- 
faced to the A/D board. Sensors are available 
to cover a wide range of pressures. The sensors 
with a range of 0 to 103.4 kPa (0 to 15 psi) 
have proved to be the most useful. The work 
reported in this paper used 185PC15DT sen- 
sors from Micro Switch (Freeport, IL) that 
require an excitation voltage of 7 to 16 V and 
give a signal of 0 to 5 V on a 1-V baseline. 
The sensors are designed for printed circuit 
board mounting. A small wooden block cany- 
ing a three-conductor audio jack (274-249A; 
Radio Shack, Fort Worth, TX) was cemented 
to the back of the sensor, and connections were 
soldered from the three sensor leads (excitation 
in, signal out, and ground) to this jack to allow 
secure andeasy electrical connection or 
removal of the sensor. Sensors were connected 
via Viton tubing (Number 64 12- 16; Cole- 
Palmer, Chicago, IL) to a Luer adaptor (Num- 
ber 6155; Popper & Sons, New Hyde Park, 
NY). Hose connections were secured with ny- 
lon clamps (Number L-06832-03; Cole- 
Palmer). Metal hub needles (20 gauge, 2.54 
cm; Number 7056; Popper & Sons) were 
cemented with epoxy glue to the Luer adapters 
to provide leakproof junctions. When neces- 
sary, needles were replaced by soaking the 
adaptor and needle combination in acetone to 
loosen the glue bond. 
MD Intelface Board and Computer. Most 
of the data were obtained with the Real Time 
Devices (State College, PA) AD2000 interface 
board. This board can read 8 differential or 16 
single-ended inputs and contains a programma- 
ble amplifier for direct use with either differen- 
tial or single-ended sensors. Electrical connec- 
tions to the interface board were made via the 
TB40 terminal board from Real Time Devices. 
More recent studies employed an AD2200 
board from the same source. This board uses a 
50- rather than 40-pin connector, which is 
advantageous when more than 16 input lines 
are desired. If the AD2200 interface board is 
used, a TB50 terminal board is required. Either 
80286 or 80386 IBM@-compatible computers 
have been used. 
Sofhyare. We have used the Atlantis@ data 
acquisition software package from Real Time 
Devices. This software was designed for use 
with the Real Time Devices boards and sup- 
ports the programmable amplifier feature of 
the boards. The current version of the software 
reads only 8 input signals, but a 16-channel 
version that runs under Windows Version 3.0 
or 3.1 (Microsoft, Redmond, WA) is now 
available. 
Methods 
Substrate and Inoculum Preparation. The 
forage was ground through a 1-mm screen in a 
Wiley mill (Arthur H. Thomas Co., Philadel- 
phia, PA) after drying in a 60°C oven. The 
alfalfa hay used in the experiments on incuba- 
tion conditions and calibration contained 
49.4% NDF (ash-free, DM basis), 38.9% ADF, 
and 8.1% lignin. 
The ruminal fluid inoculum used in all ex- 
periments was obtained from a nonlactating 
Holstein cow that had ad libitum access to 
medium quality mixed mostly grass hay. She 
was fed twice daily. The ruminal fluid was 
filtered through four layers of cheesecloth and 
then through glass wool before it was added to 
the in vitro medium. Normally, ruminal fluid 
was collected 2 h after feeding, but in some 
cases collection time varied by as much as 3 h. 
Incubation Conditions. The phosphate- 
bicarbonate medium and reducing solution of 
Goering and Van Soest (5 ) were used for all 
incubations. Forage (100 mg) was transferred 
to a 50-ml serum bottle and wetted with 1.0 ml 
of boiled distilled water that had been cooled 
to room temperature. The medium was heated 
separately to just below boiling under C02 and 
Journal of Dairy Science Vol. 76, No. 4, 1993 
1066 PELL AND SCHOFIELD 
then cooled in ice. Reducing solution (.4 ml) 
was added to each serum bottle, followed by 
addition of 6.6 ml of medium. Strict anaerobic 
technique was employed in all transfers (1, 7). 
The bottles were stoppered with previously 
unused, lightly greased butyl rubber stoppers 
(GeoMagnetic Technologies, Ochelata, OK) 
and crimp sealed. After the medium was 
equilibrated for 5 min in the incubator, ruminal 
fluid (2.0 ml) that had been filtered through 
glass wool was added by injection, and the 
pressure sensors were inserted. After a further 
15-min equilibration at 39'C, the pressure in 
each bottle was zeroed by puncturing the s top 
per with a needle for 5 s. The minimum time 
for equilibration was not established, and an 
interval of less than 15 min possibly could be 
used. The sensors then were plugged into the 
computer leads, and readings were initiated. 
The usual interval between readings was 60 
min, and each reading was an average of the 
sensor voltage taken over a 204 interval. Data 
were displayed on the monitor and were auto- 
matically written to the hard disk as an ASCII 
file that was imported into a spreadsheet pro- 
gram for further processing. The calculations 
to correct for the blank values and for conver- 
sion of the net voltage into gas volume using 
the appropriate calibration constant were made 
using a spreadsheet macro. All gas volumes 
were corrected to reflect DM input. 
For the experiment to determine appropriate 
inoculum size, clarified ruminal fluid was 
needed as an ingredient in the medium. This 
fluid was clarified by centrifugation at 26,890 
x g under C02 for 15 min at 20°C. 
Calibration of Sensor Output. Four 50-ml 
serum bottles containing 10 ml of incubation 
medium (prepared exactly as for an in vitro 
incubation) were equilibrated at 39'C, pressure 
sensors were inserted, and the pressure was 
zeroed by needle puncture. Several 125-ml se- 
rum bottles were filled with CO;! to a pressure 
of about 68.9 Wa, and these bottles were 
equilibrated in the incubator. Aliquots of gas 
(3 ml) were injected into the stirred 50-ml 
bottles, and a pressure reading was taken after 
each addition. Sufficient time was allowed for 
the reading to stabilize before the next gas 
aliquot was added. A similar procedure was 
followed using air instead of C02 as the added 
gas to generate the response curve for a gas 
with low water solubility. 
Journal of Dairy Science Vol. 76. No. 4, 1993 
The sensors used give a differential pressure 
reading: the difference between atmospheric 
pressure and the pressure at the sensor input. 
Because all pressure values used for volume 
calculations are themselves differences be- 
tween sample and blank, changes in the exter- 
nal atmospheric pressure have no effect on the 
volume determination. 
Measurement of Headspace Gas Composi- 
tion. As an alternative to gas chromatography, 
the fraction of CO;! in the headspace gas was 
measured by treating the gas mixture with 
NaOH solution and observing the pressure 
drop. The method to measure the composition 
of the headspace gas is a modification of the 
approach used by Hungate et al. (9). Serum 
bottles (10 ml) containing magnetic stir bars 
and 5 ml of 10% NaOH at 39'C were injected 
with 10 ml of air, and the pressure change was 
recorded. The pressure then was reset to at- 
mospheric level by needle puncture, and 10 ml 
of headspace gas were injected. After stirring 
and reequilibration, the final pressure was 
recorded. The ratio of these two pressure read- 
ings (mixture:air) gives the fraction of insolu- 
ble gas (mostly CH4) in the mixture. 
Micro-NDF Analysis. A 100-mg forage 
sample was suspended in 10 ml of in vitro 
digestion buffer in a 50-1111 serum bottle, and 
20 ml of the standard NDF reagent ( 5 , 21) 
were added. The bottle was capped, crimp 
sealed, and autoclaved at 105°C for 60 min. 
The contents were filtered hot through 
preweighed glass fiber filters (Gelman A/E, 47 
mm; Gelman Sciences, Ann Arbor, MI), 
washed three times with 50 ml of hot water, 
rinsed with ethanol and acetone, and dried in 
aluminum weigh pans for either 2 h in a 
vacuum oven at 85°C or overnight at 100°C. 
These two drying methods gave similar results. 
The pans were allowed to cool for 1 min in a 
closed chamber containing phosphorous pen- 
toxide before they were weighed. The general 
linear models procedure of Minitab (13) was 
used to test differences between the micro- 
NDF method and that of Van Soest et al. (21). 
This micro-NDF method has not been vali- 
dated for use with concentrates. 
Preparation of NDF Forage Extracts for 
Digestion Trials. Triplicate 400-mg samples of 
alfalfa (NDF. 43.8%), timothy (NDF, 72.0%), 
and corn stover (NDF, 73.3%) were extracted 
with standard NDF solution for 1 h at 105°C in 
COMPUTERIZED MONITORING OF GAS PRODUCTION1067 
the autoclave. Termamyl amylase (50 p1; 
Number TC 1000-2436; Fisher Scientific, 
Pittsburgh, PA) was added to the alfalfa sam- 
ple (21). For each forage, the samples were 
pooled, filtered on a sintered glass funnel, 
washed five times with 100 ml of hot distilled 
water, and allowed to soak for 30 min in 1 M 
magnesium acetate to remove final traces of 
detergent. After four more hot water washes, 
the forage was rinsed with ethanol and acetone 
and dried overnight at approximately 70°C in a 
vacuum oven. 
RESULTS AND DISCUSSION 
Calibration Curves and Gas Composition 
Because in vitro digestion produces a mix- 
ture of a water-soluble gas (C02) and an es- 
sentially water-insoluble gas (CHd), the cali- 
bration must measure the sensor response 
curves to both types of gas. The sensors 
showed the expected linearly increasing volt- 
age response to increasing amounts of air and 
C02 (Table 1). The reciprocal slopes of these 
calibration curves were 8.88 m W for C02 
(calculated value, 8.68 m W ) and 7.66 m W 
for air (calculated value, 7.68 mW). The A p 
pendix contains more extensive details and 
calculations for the calibrations for soluble and 
insoluble gases. The relatively high water solu- 
bility of C02 causes the slope of the plot of 
volume versus voltage to be greater than that 
for a low solubility gas. 
For both air and C&, the agreement be- 
tween the calculated and the observed volume: 
voltage ratios shows that the sensors are per- 
forming linearly and to specification. For a 
mixture of soluble and insoluble gas, the cor- 
responding voltage to volume conversion fac- 
tor (C) can be calculated from the formula: 
c = ci x C$(f x ci + (1 - f ) x C,) 
where Ci and C, are the conversion factors for 
air and C02, and f is the fraction of C02 in the 
headspace gas. The range of likely f is .9 to .6 
(8, 24), corresponding to conversion factors of 
8.74 to 8.36. Thus, even the most extreme shift 
in methane (or hydrogen) output causes an 
approximate shift of only 4% in this factor. 
For the most accurate work, a determination of 
the amount of C02 in the headspace gas 
TABLE 1 . Calibration of sensors. Known volumes of 
either air or Co;? were injected into 5 0 4 serum bottles 
that contained 10 ml of C02-saturated in vitro medium, 
and pressure changes were measured. 
Gas Readings Range Slope R2 
(no.) (W) 
Air 6 2-12 7.66 ,9993 
co2 13 1-35 8.88 ,9998 
clearly is advisable. For many purposes, how- 
ever, the use of a standard of .85 suffices, and 
this value was used to convert voltage changes 
into gas volumes at 39'C. 
Amount of Forage and Vessel Size 
Approximately 45 ml of gas was produced 
in the fermentation of 200 mg of grass hay 
during 24 h (12). This volume produces a 
pressure increase of about 103.4 kPa in an 
incubation volume of 45 ml, which cor- 
responds closely to the volume available in a 
Wheaton 50-ml serum bottle (Wheaton Inc., 
Millville, NJ) containing 10 ml of liquid. To 
avoid operation of the sensors at close to their 
maximum reading, a standard forage input of 
100 mg and serum bottles of 50 ml were 
chosen. This input is 20% of the amount typi- 
cally used in an in vitro digestion, and sam- 
pling errors become more significant. These 
errors can be controlled by appropriate dupli- 
cates. 
Because vessel size and available gas vol- 
ume can be changed readily, this pressure- 
based system is much more flexible than one 
based on volume measurements. We have ana- 
lyzed as little as 20 mg of plant cell walls by 
reducing the volume of the serum bottle. The 
incubation volume and pressure sensor range 
also can be varied to adjust the sensitivity of 
the system. The limiting factor in such scaled 
down experiments is the contribution of the 
blank that plays a more important role when 
the forage input is decreased. Under standard 
conditions, blanks are 15 to 20% of the sample 
after 24 h. For comparison, the system of 
Menke and Steingass (12). which uses a larger 
relative input of ruminal fluid, gives 24-h 
blanks of about 30%. 
Journal of Dairy Science Vol. 76, No. 4, 1993 
1068 PELL AND SCHOFIELD 
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Figure 2. Effect of increasing the size of the ruminal 
fluid inoculum on gas production of 100 mg of alfalfa. A) 
Unclarified ruminal fluid was used as the inoculum: .25 ml 
(+), .5 ml (a), 1.0 ml (A), and 2.0 ml (V). B) .5 ml (a), 
1.0 ml (A), 2.0 ml (V), and 4.0 ml (*) of unclarified 
ruminal fluid were used as the inoculum with sufficient 
clarified ruminal fluid to bring the total amount of ruminal 
fluid to 4 ml. 
Volume of Medium and Ruminal 
Fluid Inoculum 
The amount of liquid added to the 
100-mg forage samples was set at 10 ml to 
maintain the same 1iquid:sample ratio as used 
in the protocol of Goering and Van Soest (5) . 
Because blanks are related directly to inocu- 
lum volume, a small inoculum is advanta- 
geous, provided that gas production is not 
limited by inoculum size. In Figure 2A, the 
results of digestion of a fixed amount (100 mg) 
of alfalfa with increasing amounts of ruminal 
fluid suggest that a 20% inoculum is sufficient 
to ensure the maximum rate of fiber digestion 
but that lower percentages are not. This finding 
supports the recommendations of Goering and 
Van Soest ( 5 ) for in vitro digestion. To deter- 
mine whether the differences in rate were due 
to the number of microbial cells in the inocu- 
lum or to some factor in the ruminal fluid, a 
second experiment was conducted. Clarified 
ruminal fluid was added to the tubes so that 
the total amount of ruminal fluid, both clari- 
fied and unclarified, was similar among tubes. 
The smaller inocula appeared to have a slightly 
longer lag time than larger inocula, but the 
maximum gas productions were similar 
(Figure 2B). The lag period reflected the time 
required for the microbial numbers to increase 
to levels comparable with those in the larger 
inocula. The difference in results between the 
two experiments with and without added clari- 
fied ruminal fluid suggests that a factor in the 
clarified ruminal fluid affected microbial 
growth. 
Another important consideration is whether 
or not the ruminal fluid is homogenized in a 
blender before glass wool filtration. Although 
blending may increase the number of previ- 
ously fiber-adherent bacteria in the inoculum, 
it also increases the number of small feed 
particles. As a result, when the samples were 
blended, blanks were higher than values for 
blended samples. Most of the inocula used in 
these experiments were prepared without 
blending, but early experiments in which 
blending was used gave results similar to those 
of the Vials with the unblended inocula. In our 
laboratory, inocula are not blended because 
blending increased the gas production of the 
blanks, introduced another procedure in the 
protocol that could expose the microbes to 
oxygen, and results were similar with and 
without blending. 
Effects of Stirring 
Eight identical corn silage samples, four 
with stirrer bars and four without, were in- 
cubated for 42 h under standard conditions. 
The plots of gas volume versus time are shown 
in Figure 3. Individual data points were omit- 
ted from these plots to reveal differences bet- 
ter. The coefficient of variation among the 
maximum readings was 2.1% for the stirred 
Journal of Dairy Science Vol. 76, No. 4, 1993 
COMPUTERIZED MONITORING OF GAS PRODUCTION 1069 
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Figure 3. The effect of stirringon the within run 
reproducibility. Eight 1Wmg samples of the same corn 
silage were monitored: four were stirred intermittently, 
and four were unstirred. A) Stirred, B) unstirred. 
samples and 4.2% for the unstirred samples. 
Consequently, stirring was used for all subse- 
quent analyses. Stirring may be important be- 
cause COz has a strong tendency to form 
supersaturated solutions, and this tendency is 
minimized in the presence of a finely divided, 
stirred, solid suspension. 
In some cases, the forage tended to adhere 
to the vessel walls. To minimize this interfer- 
ence, the serum bottles were disconnected 
from the sensors and swirled by hand to wash 
down adherent fiber. These hand rotations 
were performed two or three times during a 
48-h run. 
20 I 7 
31 5 
2 10 - 
0 5 10 15 20 25 30 35 40 45 50 
Hours 
Figure 4. Coefficients of variation within run for six 
samples of alfalfa using the same inoculum. 
Reproducibility 
This parameter was tested in two ways: 1) 
within a series of samples incubated with ali- 
quots of the same ruminal fluid and 2) among 
different series in which the ruminal fluid sam- 
ple was taken at the same time (2 h after the 
morning feeding) but on different days. The 
same alfalfa forage was used in all these trials. 
The reproducibility within run is demonstrated 
in Figure 4, in which the coefficients of varia- 
tion for six identical alfalfa samples, all weigh- 
ing 100 mg, were plotted as a function of time. 
Except for the first time point (at which the 
volume of gas produced was less than 1 ml), 
the coefficient of variation was less than 4%. 
The reproducibility among different runs 
(Figure 5 ) is similar in pattern but varies more 
in the first 9 h and reaches an essentially 
constant value of about 4% after 12 h. In some 
early experiments that are included in this 
analysis, the time of collection of ruminal fluid 
varied by as much as 3 h from the standard 
2-h postfeeding schedule later adopted. 
Reproducibility among runs may be improved 
when a strict ruminal fluid collection schedule 
is used. 
Validation of Micro-NDF Method 
Weimer et al. (22) used a modification of 
the standard NDF method (5 , 21) for analysis 
of the digestion kinetics of various pure cellu- 
Journal of Dairy Science Vol. 76, No. 4, 1993 
PELL AND SCHOFIELD 1070 
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h 8 15 
2 10 
Y 
5 
0 
I 
'iz 
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Figure 5. Hourly coefficients of variation from 13 
experiments using different inocula and alfalfa hay as the 
substrate. 
lose fractions by ruminal bacteria. The neutral 
detergent was used simply to solubilize the 
microbial component of the mixture. In the 
Weimer method (22), neutral detergent solu- 
tion was added to the in vitro incubation mix- 
ture at the end of the digestion, and the sealed 
vessel then was autoclaved for 45 min at 
125'C. If this procedure is applied without 
modification to forage samples, the NDF ob- 
tained are somewhat lower than those found 
using the standard method (Pel1 and Schofield, 
1992, unpublished data). If, however, the au- 
toclave conditions are modified to 60 min at 
105'C, NDF for a variety of forages cor- 
respond closely to those obtained using the 
standard method (Table 2). Neither the differ- 
ences between the two methods (P > .68) nor 
the forage by method interaction was signifi- 
cant (P > .30). The modified NDF method 
allows analyses of samples of 10 to 50 mg of 
residual fiber, and the method is convenient 
and rapid. The small sample size precludes 
correction for ash. 
Correlation of Gas Production 
with NDF Disappearance 
In an intact forage, gas produced during in 
vitro digestion comes from both the soluble 
and the fiber fractions. To correlate gas 
production with NDF disappearance, samples 
of alfalfa, corn stover, and timothy were ex- 
tracted with NDF solution to remove the solu- 
ble fraction. Increasing amounts of the washed 
NDF residue of each forage then were 
digested, gas production after 48 h was deter- 
mined, and the residual NDF also was meas- 
ured using the previously described microas- 
say. Figure 6 shows the digestion curves for 
the three forages. Although the overall patterns 
were similar, digestion of alfalfa clearly termi- 
nated much earlier than that of the corn stover 
or timothy, possibly because of the higher 
lignin content of the former (7.8% for the 
legume compared with 6.5% for the timothy 
and 4.0% for the corn stover). 
The relationship between gas production 
and NDF consumption is shown in Figure 7. 
The linear correlation (R2 = .99; n = 15) 
suggests that essentially the same chemical 
reactions occurred in the digestion of each 
of these three forage NDF preparations. 
The regression equation that best fitted 
the data was Y = 1.415 + .346 X where Y = 
gas (milliliters) and X = NDF disappearance 
TABLE 2. Comparison of micro-NDF and standard NDF techniques. The NDF values for both methods are based on 
three replicates. Least squares means and standard errors of the mean are presented. 
NDF (micr0)l N D F 
- 
Forage Tz SEM cv X SEM cv 
Alfalfa 18.95 .52 2.32 20.06 .52 1.57 
Bromegrass 43.01 .52 1.13 42.58 .37 1.04 
Alfalfa 49.33 .52 1.68 48.77 .52 .46 
Timothy 71.48 .52 .42 7 1.63 .37 .81 
Stargrass 86.03 .52 1.28 85.12 .37 1.30 
'Determined by heating in autoclave for 1 h at 105'C for 60 min in detergent solution (21) and filtering. 
2According to method of Van Soest et al. (21). 
Journal of Dairy Science Vol. 76, No. 4, 1993 
COMPUTERIZED MONITORING OF GAS PRODUCTION 1071 
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v 
In 
d 10 
5 
0 
A I 
0 5 10 15 20 25 30 35 40 45 
Hours 
30 
B 
25 
20 
E 
T 1 5 
10 
5 
0 
3 
0 5 10 15 20 25 30 35 40 45 
Hours 
35 
30 
25 
E 20 
rn d 15 
10 
h 
Y 
C 
5 
0 
0 5 10 15 20 25 30 35 40 45 
Hours 
Figure 6. Gas production of increasing amounts of 
forage: 25 mg 50 mg (e), 75 mg (A), 100 mg (V), 
and 125 mg (+). Alfalfa (A), timothy (B), and corn stover 
(C) NDF residues were used as the substrate with a fixed 
volume (2 ml) of ruminal fluid. 
0 4 a 3 r r , 0 1 0 2 0 3 0 4 0 5 0 6 0 7 0 8 0 
NDF Disappearance (me) 
Figure 7. Correlation of gas production with decrease 
in NDF (milligrams). Decrease in NDF was determined by 
measuring the initial and residual NDF of alfalfa (A), corn 
stover (0). and timothy (m) when 50, 75, 100, 125, 150, or 
175 mg were used as the initial substrate with 2 ml of 
ruminal fluid. 
(milligrams). When the y-intercept was as- 
signed a value of zero, and the line was forced 
through the origin, the x-coefficient was .373, 
and the R2 was .W. 
Despite that correlation, adjustments must 
be made when feeds that produce different 
VFA profiles are compared. When forages are 
fermented, the differences in the amount of 
propionate produced are small (12), but, when 
concentrates are digested, the acetate to propi- 
onate (A:P) ratio and the amount of C02 
produced decrease. Menke and Steingass (12) 
addressed this question by developing three 
separate equations for concentrates, forages, 
and mixed feeds to convert gas production to 
metabolizable energy. Work is underway in 
our laboratory to measure the A:P ratio over 
time with different substrates and to correlate 
these values with gas production, gas composi- 
tion, and digestion. 
In addition to its production during fermen- 
tation, C 0 2 evolves from the buffer when VFA 
are neutralized. Gas evolution from the rumi- 
nal fluid inoculum is accounted for by blanks. 
A buffer that maintains a constant pH is neces- 
sary, or gas evolution will become pH- 
dependent. Preliminary data from our labora- 
tory and work by Grant and Mertens(6) have 
confirmed that the buffer described by Goering 
and Van Soest (5 ) maintains pH during the 
course of an incubation with forages. Because 
Journal of Dairy Science Vol. 76, No. 4, 1993 
1072 PELL AND SCHOFIELD 
this buffer contains high concentrations of 
phosphate, the molar yield of C 0 2 from the 
reaction of VFA with bicarbonate is <1. Gas 
production data would be easier to interpret if 
a buffer system based entirely on either bicar- 
bonate or phosphate could be used. However, 
to meet microbial requirements for bicarbonate 
(2) and pH > 6 (14), a mixed bicarbonate- 
phosphate buffer system is necessary. When 
high concentrate samples are digested, pH 
measurement is recommended. 
CONCLUSIONS 
Analyses of gas production and change in 
NDF measure different processes. Traditional 
in vitro methods follow disappearance of one 
component of the substrate, whereas gas meas- 
urement focuses on the appearance of fermen- 
tation products. These products are the result 
of the fermentation of both soluble and insolu- 
ble substrates, which is an advantage over the 
traditional method that does not consider the 
soluble substrates. Digestion rates calculated 
using the gas production measurements reflect 
multiple rates from the soluble and insoluble 
fractions. Under conditions that are not 
nutrient-limiting, gas production is a direct 
measure of microbial growth and, in some 
respects, is a better index of forage metaboliza- 
ble energy yield than is the indirect measure 
based on NDF decrease. Analyses of gas 
production consider all of the wealth of major 
metabolic energy sources and measure those 
different sources (monosaccharides, polysac- 
charides, pectins, starch, cellulose, and 
hemicellulose) in the same currency, namely, 
their conversion to C 0 2 and CH4. This method 
can be used to determine the importance of 
some of these different feed fractions in 
providing energy to the microbes and to deter- 
mine whether compounds inhibit microbial ac- 
tivity. The small sample size makes this tech- 
nique especially useful with tissue culture 
samples or synthetic compounds. 
ACKNOWLEDGMENTS 
The support of the Fats and Protein Re- 
search Foundation in providing some of the 
funding for this research is greatly appreciated. 
Also, the authors thank T. Hernandez for his 
assistance in analyzing the forage samples. 
REFERENCES 
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APPENDIX 
Differences in Sensor Response 
to C o p and CH4 
Insoluble Gas. In the calibrations for insolu- 
ble gas, the available gas volume was 48 ml in 
a Wheaton “50-ml” serum bottle with 10 ml of 
liquid. 
If added gas volume = X ml, 
then pressure increase = W48 am, V I 
and voltage change is 5 x 10/8 = 6.25 V/atm [2] 
(for a sensor calibrated to produce a 5-V/atm 
response at an excitation level of 8 V and an 
actual excitation of 10 V). Thus, X ml of gas 
produces a voltage change of 6.25 x XI48 V, 
and gas volume = 7.68 x net voltage change. 
When air was used for calibration, an excellent 
linear response was found with a volume: 
voltage ratio of 7.66 (Table 1). 
Soluble Gus. Henry’s Law gives the rela- 
tionship between gas pressure and solubility. 
For C O 2 at 37°C this relationship takes the 
form 
[C02laq = ,0246 x P 
where dissolved C 0 2 is in moles per liter, and 
the pressure is in atmospheres (3). From the 
universal gas law, 
[C02lgas = P/RT 
where R = .082 L x atd(mo1e x OK). Thus, 
[C02]&[C02]g.s = .0246 x RT. 
If we assume a liquid volume of 10 ml and a 
gas volume of 48 ml, then volume 
(C02)&(C02)gas = .0246 x RT x 10/48 
= .130 at 37°C. 
Thus, if 10 ml of C O 2 are added, 8.85 ml will 
remain available to cause a pressure change. 
From Equations [ l ] and [2], 
voltage change = 6.25 x 8.85/48 V = 1.152 V 
andmilliliters of gasN = 1011.152 = 8.68. 
Journal of Dairy Science Vol. 76, No. 4, 1993