Field-flow fractionation of alkali-liberated nuclear polyhedrosis virus from gypsy moth Lymantria dispar linnaeus

Journal of VirologicalMethods, 0 Elsevier/North-Holland

FIELD-FLOW

Biomedical

241

241~ 256

Press

FRACTIONATION

OF ALKALI-LIBERATED

NUCLEAR

VIRUS FROM GYPSY MOTH L YMANTRZA DZSPAR LINNAEUS

POLYHEDROSIS

KARIN

1 (1980)

D. CALDWELL’,

THANH

T. NGUYEN’,

J. CALVIN

GIDDINGS’

and HORACE

M.

MAZZONE’ r Department

of Chemistry,

of Agriculture, (Accepted

6 March

Rod-shaped through ated

contains

FFF)

molecular

a wide mixture

weights

it was possible

the same viral mixture technique.

FFF

effective

molecular

scopic

which

of the enveloped

weight

and ’ U.S. Department U.S.A.

confirmed were material

thinner

virus from

of enveloped

aggregated

by means

of sedimentation resolved

forms

commonly

against

population

that

this lower the initial

had been removed

molecular

weight

enveloped

pattern

Through

electron

dimers,

In parallel

gradient

sedi-

of manipulation,

from each separation. detergent

of non-enveloped fraction

etc.

used density

a nonionic

monomers

fractionation

the elution

components.

was lower than that of the initial monomer

than

field-flow

as monomers,

The liber-

as well as enveloped

From

that can be extracted

was dialyzed

were obtained hosts.

as to time and convenience

information a uniform

(NPV)

infected

particles.

by the more

are compared

revealed

which

polyhedrosis

the components

viral structures

of the dialysate

observations

structures

UT 84112; CT 06514,

and purified

to the various

was separated

methods

nuclear

of the separated

be assigned

of physicochemical

of aggregate

mentation

spectrum

to characterize

The two

as well as to the amount The mixture

City,

Hamden,

isolated

was separated

could

experiments

moth

bodies

assortment

to give a mass

microsmcopy mentalion

Station,

of the gypsy of inclusion

This complex

(sedimentation effective

particles

dissolution

fraction

monomers.

of Utah, Salt Lake

Forest Experiment

1980)

viral

alkaline

viral

University

Northeastern

fraction. did indeed

indicating

solution. particles

that

Sedi-

with

Electron

an

micro-

contain

rod-like

most

if not all

from the viral rod structures.

INTRODUCTION

In the

search

for

non-chemical

agents

to be used

as biological

pesticides,

interest

has

focused on the baculoviruses with their seemingly well established host specificities (Ignoffo, 1968). These rod-shaped virus particles are known to assemble in the infected organism into large protein embedded clusters called inclusion bodies. These clusters are generally chemically inert but dissolve upon exposure to the alkaline pH characteristic of the intestinal tract of the host, thus releasing the viral rods as a mixture of enveloped monomers and higher order aggregates (Summers and Volkman, 1976; Summers and Smith 1978; Harrap et al., 1977; Rye1 and Cline, 1971). These aggregates are not artifacts due to the liberation procedure as electron micrographs of intact inclusion bodies show different sized bundles or viral rods already present within the protein matrix

242

(Ignoffo,

1968; Lee and Miller, 1979; Mazzone et al., 1980). Purification

is most commonly trifugation mentary)

carried out using density gradient

techniques. approach

flow fractionation

sedimentation

We wish to present here an alternate

to this problem (sedimentation

of these lysates

and differential

cen-

(and in some ways comple-

that is gentle and relatively fast: sedimentation

field-

FFF).

THEORY

Field-flow

fractionation

(FFF)

is a class of one-phase

separation

techniques

in which

injected samples are compressed into layers varying in mean thickness depending on their interaction with a given field acting perpendicularly to the direction of flow through a narrow ribbon-like channel. The laminar flow carries the various solute zones along at velocities which uniquely depend on their layer thickness R (Giddings, 1976; Giddings et al., 1975, 1980). The polarizing effect of the field is counteracted by diffusion and the resulting steady-state distribution of the solute is determined by the ratio of the species diffusion coefficient D to the field induced velocity U; II = D/U. More important than Q is the dimensionless layer thickness A, which is the ratio of II to the thickness w of the flow channel. We have

Rentention

ratio R in FFF can be expressed in terms of X as follows:

R = 6X[coth (1/2X) - 2h] The parameter

(2)

R is the ratio of the migration

velocity of a given retained

of an inert component and is measured experimentally V” to the elution volume of the retained peak y;

R = P/K In the limit of high retention R r 6h

peak to that

as the ratio of the channel volume

(3) (small R values) Eq. 2 takes the simple form: (4)

Depending on the nature of the imposed field, h and thus R will reflect the particular property of the sample which makes it susceptible to the field. In the case of sedimentation FFF, A, can be expressed as (Giddings et al. 1974): A,=

kT g. m Wp,~ w

(5)

243

where g is the gravitational is p, ,Ap the difference the Boltzmann in revolutions

g=( 3gI

acceleration,

m the mass of the sample particle whose density

in density between

constant.

Acceleration

solvent and solute, T the temperature

g is determined

from knowledge

and k

of the spin rate

per minute (r.p.m.) and the channel curvature (radius I):

. Xn)2

.

r

Of the factors governing h,, the temperature, spin rate, channel radius, and channel thickness are selected by the operator or instrument designer. The sample’s specific contribution to h, is the effective mass (actual mass less buoyant mass) m’ of the particle: m’ = m . rip/p,, In molar terms, Eq. 7 can be written as

m’. N:=M’

=M.

ApIp,

where M is the molecular

(8) weight of the sample and N is Avogadro’s number.

In the ab-

sence of a good value for the solute density we can evaluate the effective molecular weight M’ more definitely than the absolute value M. Cast in terms of M’, h, becomes x, = M.

RT

(9)

g.w

The combination

of Eqs. 3, 4 and 9 show that elution

tion of M’ at high retention

volume

V, becomes a linear func-

levels, which means that the fractogram is essentially

a linear

mass spectrum of the separated particles of a mixture. MATERIALSANDMETHODS

Preparation of viral inclusion bodies Inclusion bodies were isolated and purified by procedures already described (Mazzone et al., 1970; Breillatt et al., 1972). Briefly, the procedure used in this study was as follows. Larvae of gypsy moth (Lymantrzizdispar Linnaeus) were infected by the addition of stock viral inclusion bodies to their diet. Infected larvae were collected and homogenized in water. The suspension was strained through cheesecloth to remove debris and centrifuged at low speed to remove large contaminants. The supernatant was then layered onto a sucrose gradient and centrifuged at 30,000 r.p.m. for 3 h in a K 10 zonal rotor. The inclusion bodies which banded at an average density of 1.2690 (57% sucrose at 20°C) were recovered, freed of sucrose by sedimentation, ritxed several times in water, air dried and stored at 4°C).

244

Lysis procedure and isolation of the rod fraction The procedure for the integrity results reported

for alkaline dissolution

of the inclusion

bodies is of utmost importance

of the molecular weight spectrum of the resulting viral preparation. The below have all been obtained using a 10 min exposure of the inclusion

bodies to a 27°C lysing solution containing 0.008 M Na,COs and 0.05 M NaCl. A 20 ml freshly prepared lysing solution was used for each 100 mg portion of inclusion bodies. The pH, monitored occasionally during lysis, usually declined from the value 10.6 of the fresh solution to 9.8 after 10 min lysis. The suspension was then given a 5 min spin at 5000 r.p.m. in a Beckman Spinco Ultracentrifuge using a faed angle rotor at 4°C. The supernatant was rapidly poured through cheesecloth to eliminate debris and unsolubilized material and was then spun for 30 min at 36,000 r.p.m. The resulting pellet was freed from its alkaline supernatant (approximately 1 h after the initial exposure) and resuspended in distilled water. After a second pelleting under identical conditions to the first the virus was again suspended in distilled water (around 3 ml final volume per 100 mg treated inclusion bodies) and given a 5 min spin at 5000 r.p.m. to eliminate non-suspended material. Any subsequent characterization of the resulting material had to be carried out within at most 48 h (preferably 24 h). After this time the decay of the sample had progressed beyond the point of meaningful observation. Sedimentation FFF apparatus and operation The separation channel is 83.3 cm long, 2.0 cm wide and 254 pm thick with 4.2 ml void volume ( W). The channel is curved to fit the inside of an aluminum rotor basket and has an average radius of 7.7 cm. The basket is rotating around a vertical axis having liquid inlet and outlet ports positioned at top and bottom, respectively. Special o-ring seals form the transition between stationary and spinning parts. The details of the construction have been described elsewhere (Giddings et al., 1980). Spin rate is monitored electronically by means of a slotted disc fastened to the shaft which rotates between a light emitting diode and a light sensitive transistor. The carrier liquid is pumped into the system by either a Chromatronix

Model CMP-1 B pulse free pump or by a Gilson peristal-

tic pump of type Minipulse II. The shaft’s stationary exit port is hooked up to a LDC Model 1205 W monitor which is operating with a 254 nm light source and whose cell volume is 10 ~1. Injections were normally 100 ~1 (around 500 pg material). Immediately following injection the pump was turned off and the rotor was accelerated to the desired spin rate. The stop-flow period was generally 15 min in which time the sample migrates to the outer channel wall and establishes its steady-state distribution. The subsequent onset of flow marks the start of the run and the effluent from this point on was collected either in a finely graduated 50 ml buret or in a Gilson Model FC-80K fraction collector. The performance of the system was verified prior to each set of runs on a new virus prepara-

245

tion by determining sedimentation

the retention

FFF behavior

of polystyrene

latex beads of diameter 0.357 pm. The

of these beads has been studied extensively

in this labora-

tory (Giddings et al., 1978). Density gradient sedimentation A linear sucrose gradient was pumped simultaneously into two 40 ml collodion tubes. The volume of the gradient solution was 36 ml per tube and its density ranged from 1.283 g/ml (60% sucrose in 0.01 M Tris-HAc, 0.001 M EDTA, pH 7.4) to 1.127 g/ml (30% sucrose in the same buffer). The densities were determined pycnometrically. From the relationship between density and refractive index for these solutions it was possible to convert refractivity measurements, made on the fractions collected later during the elution of the gradient solution, into the corresponding densities. A 13 ml portion of the freshly prepared virus sample was carefully layered on top of the gradient solution and the tube was spun at 24,000 r.p.m. for 4 h in an SW 27 swinging bucket rotor. Fractions of 0.6 ml were subsequently collected from the bottom of the tube after passage through an LDS Model 1205 UV monitor detecting at 254 nm. Fractions containing a peak of interest were pooled, diluted six-fold and spun at 36,000 r.p.m. in a futed angle rotor for 30 min. The pelleted materials were removed from their sucrose-containing

supernatants

and resuspended

in distilled water.

Detergent dialysis One part of the initial virus suspension

was dialysed

for 18 h at room temperature

against a 0.01 M Tris-HAc, 0.001 M EDTA buffer of pH 7.4, which contained 2% (v/v) of the nonionic detergent Triton X-100 from Fisher Scientific Corp. Prior to filling, the dialysis

tubing

(Visking,

washed with the detergent

1 cm flat width)

was soaked in distilled

water and carefully

solution.

Electron microscopy Droplets of the various fractions were deposited onto formvar coated 200 mesh grids (Ernest Fullam, Inc.). One min after deposition the excess liquid was blotted off and the grids were allowed to dry. These grids were then examined in a Philips Transmission Electron Microscope (EM 201) and the sizes of observed structures were determined by comparison with pictures taken at identical magnification of a carbon grating replica with 28,800 lines per inch (Ernest Fullam, Inc.). General procedure Fig. 1 shows in schematic form the course of analysis of the alkali-liberated viral structures which was followed in this work. In accordance with this outline a freshly

246

virus containing inclusion bodies

Dialysig against nonionic detergent eliminates viral envelope

sedimentation

Sedimentation FFF of collected bands with determined density yields hydrated mass of respective fraction I

Sedimentation FFF demonstrates one single peak whose mass is lower than that of untreated monomer

I

Fig. 1. Schematic followed

Sedimentation FFF of viral suspension yields spectrum of effective mass

illustration

of the course

of analysis

of gypsy moth

nuclear

polyhedrosis

virus (NPV)

in this report.

prepared sample was split into three parts, one of which was used to yield a ‘mass spectrum’ by means of sedimentation FFF, one of which was banded out in a sucrose density gradient, and one of which was dialysed against a nonionic detergent for possible elimination of the envelope surrounding the various arrangements of nucleocapsids. In our initial studies of lysates from gypsy moth inclusion bodies we observed a number of ill-resolved peaks with the fraction containing the smallest rod-like structures corresponding to an effective molecular weight of around 130 X 106. The lysis conditions used at the time were relatively harsh, involving 30-45 min exposure to pH 10.5 at 35°C. Attempts at lysing with a 0.005 M Na,C03 solution were successful with some batches of inclusion bodies and totally ineffective with others.

247 In subsequent

min exposure

work, the conditions to a lysing solution

room temperature.

The molecular

FFF stayed relatively drastically

constant

of preparation

containing

were standardized

0.008 M NazCOs

weight distrrbution

for the &st 24-48

towards lower masses thus indicating

to involve 10

and 0.05 M NaCl at

as observed through sedimentation h after preparation,

but later changed

the decay of the sample.

RESULTS

Fig. 2 demonstrates

a typical fractogram of a freshly prepared sample of viral particles

from lysed inclusion bodies from gypsy moth larvae, &tamed as described under Materials and Methods. The first eluted peak is the void peak (elution volume VQ), which con-

i

Relaxation

EPFECTIVE

TIME

Fig. 2. Fractogram obtained through sedimentation

MOLECULAR WEIGHT

(IL

IO+’f

(minutW3)

FFF of freshly prepared alkali-liberated

Viral

rod

particles of gypsy moth NPV. Flow rate: 23.7 ml/h; spin rate: 1417 r.p.m. Shaded areas in peaks A and B represent mater% studied in the ekctroa microscope,The abeissa is graduated in time (minf, aS weli as,effective molecular weight of eiuate. Following injection, a 15 min relaxation period was observed., After the emergence of peak C, the field was turned off.

tams all components of the sample with effective mass less than 50 X IO” g/m& As the sample decays upon aging, this peak increases in size at the expense of the more massive and thus more highly retained components. The major peak, labeled A in the fqure, contained the single enveloped nucleocapsids as shown in Fig. 3. Of 100 rod-like structures, originating from the shaded zone in peak A of Fig. 2 and observed in the electron microscope, not a single agregate (dimer or larger size bundle) appeared. Peak B of the fractogram contained mostly dimers which were enveloped in such tight configuration that the two separate rods could barely be distinguished in the eiec-

248

lU

249

tron micrographs.

The length of the structure

single enveloped

rod, whereas the thickness

mer. In peak C trimers with the structure

were clearly present,

(see Fig. 4) is approximately

that of the

is more than twice the value for the monosome with visibly intact envelopes

and some

partially ruptured.

Peaks A, B and C represent the most abundant components of the sample. If the. separation procedure had been allowed to continue, a number of minor peaks would likely have been observed. In the interest of maximizing the information obtained from one sample before the onset of the decay process, while minimizing wear and tear on the equipment, the field was generally turned off after emergence of the three major components of the sample. As the rotor comes to a stop, which occurs within 30 set, the carrier liquid begins to sweep out all material remaining in the column. This remainder fraction, which. is the last peak to elute in Fig. 2, is seen in the electron microscope to contain complex aggregates from tetramers on up. The fractogram in Fig. 2 shows the monomer peak (A) to elute after 53 min under the particular set of conditions of flow rate (24 ml/h) and field (1417 r.p.m.) selected for this study. The separation process may be speeded up by either an increase in flow rate while retaining the field strength, in which case speed is achieved at the expense of resolution, or by increasing both field and flow so as to retain resolution. The latter case is particularly

attractive

in conjunction

with flow programming

(see below),

but

Inject Start flow

I

I

TIME (minutes)

Fig. 5. Fractogram

of freshly

at a spin rate of 1801

r.p.m.

prepared

alkali-liberated

viral rod sample

and a flow rate of 60 ml/h.

gence of peak A the field was turned

off and the detector

Relaxation sensitivity

of gypsy

moth

NPV obtained

time was 15 min. After cut by a factor

the emer-

4. - .-_

Fig. 3. Electron

micrograph

of a single rod-like

Fig. 4. Electron

micrograph

of an enveloped

50 kV.

particle

obtained

from peak A in Fig. 2.

viral dimer from peak B in Fig. 2. Uranyl

acetate

stained,

250 high spin rates tax the life span of the seal between the rotating and stationary parts in the apparatus and thus meet with some practical difficulties. Fig. 5 illustrates a step towards

the optimization

of separation

time and resolution.

Both flow rate (60 ml/h)

and spin rate (1801 r.p.m.) are increased with respect to those of Fig. 2, a change permitting

a reasonably

well resolved monomer

peak to elute after 28 mm instead of 53

min. This by no means represents the ultimate performance

of the equipment,

but merely

indicates the versatility inherent in the sedimentation FFF technique. As shown by Eqs. 2 and 5, parameter h and thus retention ratio R are governed by the field strength and solvent density selected by the operator as well as by the samplespecific parameters

mass and density.

The unsolvated

density

of a virus particle is diffi-

cult to obtain with good precision. Such a determination often involves fmding the density of a virus solution of a given concentration where the solvent density is well known. The concentration has to be determined either gravimetrically by evaporating a known aliquot of the solution to complete dryness, in which case all water of hydration should be eliminated, or chemically, in which case the complete structural composition of the virus particle must be known. Both procedures are laborious and error prone (Markham, 1967; Freifelder, 1970). We have chosen here to discuss the sedimentation FFF separation in terms of the effective molecular weight M’ (see Eq. 8) which is then the sole species-specific parameter responsible for retention. An approximate hydrated density is available from studies of isopycnic sedimentation. Separation in a sucrose density gradient As shown in the analysis scheme in Fig. 1, a portion of the freshly prepared alkaliliberated virus sample was spun in a sucrose gradient solution whose density ranged from 1.283 to 1.127 g/ml. Visual inspection of the banding pattern revealed three major components fractogram

and at least four bands of lesser significance. in Fig. 6 which was obtained

This was borne

by sampling from the bottom

out by the

of the gradient

tube. Densities of the collected fractions were identified refractometrically. The resulting density profile is superimposed on the UV absorption curve in Fig. 6. The least dense band A, which represented the best resolved component, density of 1.192 g/ml and was shown in the electron microscope nucleocapsid

in monomer

had an average apparent to contain the enveloped

form. Peak B, appearing at the next higher density,

was seen to contain mostly enveloped dimers. This order of banding previously for other baculoviruses (Summers and Volkman, 1976).

1.204 g/ml,

has been observed

Cross referencing of results between density gradient sedimentation and sedimentation FFF Peaks A and B from the density gradient sedimentation were pooled separately and freed from their sucrose containing medium through pelleting at 36,000 r.p.m. and subsequent resuspension in distilled water. The resulting samples were injected one at a time

251

1.28

1.21 2: F ,M P 1.20

g 3 8

1. 16

0

I

I

I

I

5

10

15

20

EFFLUENT

VOLUME

Fig. 6. Elution pattern obtained after sucrose viral rod

sample

of gypsy

spinning

at 24,000

r.p.m.

ml were

collected

every

represents

moth Elution 2 min.

the shape of the density

proceeded Their

(ml)

density

NPV. Sedimentation from

densities

1.12

gradient

sedimentation

of an alkali-liberated

time was 4 h in a SW27 swinging the bottom

were determined

of the gradient

tube.

refractometrically.

bucket

Fractions The dotted

rotor of 0.6 line

gradient.

into the sedimentation FFF apparatus. In this manner it was found that peak A from the density gradient sedimentation is identical with the monomer peak of effective molecular weight 160 X 106, which is marked A in the sedimentation FFF fractogram (Fig. 2). Peak B from the density gradient separation could likewise be identified with peak B in Fig. 2. We have no justification for assuming that the density gradient separation in Fig. 6 is truly an isopycnic equilibrium sedimentation without participation of a rate-zonal sedimentation. If the system actually arrived at equilibrium the banding densities would be identical to the hydrated densities of the particles, provided the osmotic impact of the gradient former could be ignored. Thus, while realizing that the densities determined through this procedure are not entirely accurate, we use them to calculate approximate molecular weights for monomeric and dimeric species. According to Eqs. 2, 8 and 9, component A would represent a hydrated molecular weight of 160 X lo6 divided by 0.197/1.192 which gives 968 X lo6 g/mol. The carrier, water, is assumed to have a density of 0.995 g/ml. The dimer similarly shows a molecular weight of 1540 X 106. The fact that monomer and dimer are not different multiples of one mass is possibly accounted for by the different amounts of envelope material associated with the two arrangements and by the fact that the densities used in the calculation are not reliable values.

252

Detergent dialysis It has been observed (Summers exposure

to nonionic

baculoviruses

after

detergents

and Volkman,

1976; Summers and Smith, 1978) that

tends to eliminate

alkali liberation

the envelope

from their inclusion

associated

with most

bodies. Harrap et al. (1977)

demonstrated that the multiplicity of bands seen upon density gradient sedimentation of alkali-liberated viruses from Spodoptera littoralis can be abolished if the sample is pretreated with detergent. A fresh preparation of the gypsy moth virus suspension, consisting of a mixture of aggregate forms, was dialysed against a 2% Triton X-100 solution for 18 h and the resulting sample injected into the sedimentation FFF apparatus (see Fig. 7). As before, the carrier liquid was distilled water. A pronounced

void peak appeared and was largely

attributed to the detergent which absorbs at the 2.54 nm wavelength used for detection. The void peak was followed by a component whose mass was much lower than what was previously observed for the enveloped monomer (see Fig. 2). At its usual elution position (corresponding to an effective molecular weight of 160 X 106) there was no peak, and upon the turn off of the field only a minor amount of material eluted. A sample from the retained peak in Fig. 7 was examined by electron microscopy (see

EFFECTIVE

0

8 50 I 100

MOLECULAR

(x 10-b)

1

I

I

0

WEIGHT

200 .

150

60

30 TIME (minutes)

Fig. 7. Fractogram NPV dialysed The relaxation cular weight

obtained

against

from

a nonionic

a freshly

prepared

detergent

for

time was 15 min. The abscissa of the eluate.

alkali-liberated

18 h. Spin rate:

is graduated

viral rod sample 1403

r.p.m.;

of gypsy

flow rate:

moth

23.6 ml/h.

in time (min) as well as in effective

mole-

253

Fig. 8. Electron micrograph of detergent treated viral rods, as they emerge from the sedimentation FFF experiment depicted in Fig. 7. Elution time for this sample was 27 min.

Fig. 8):

iS rod-like

330 X 40 mn.‘The

structures standard

observed deviation

in one picture

yielded

average dimensions

of

in the length was 3%. The nature of the fracto-

gram, backed by electron microscopic observations, thus suggests that the detergent exposure causes the viruses, monomers as well as multimers, to shed their envelopes to yield a uniform

population

of monometer

nucleocapsids

whose effective molecular weight aver-

ages 78.3 X 106. DISCUSSION

The sedimentation FFF technique is an analytical separation method capable of yielding a high resolution of complex samples in relatively short times. A comparison between the two separation schemes executed in this study, sedimentation FFF and density gradient sedimentation, can be summarized in the following four points. 1. Time of separation. A clean population of enveloped nucleocapsids can be obtained in 30 min by sedimentation FFF. The density gradient sedimentation protocol described

254

above requires 4 h for the resolution of the components within the gradient solution, another hour for elution, and a final 30 min for elimination of the gradient former through pelleting and resuspension in a sucrose free medium. If the sedimentation FFF separation were allowed to run its full course to yield fractions with masses much higher than those observed here, fractionation

times could be-

come S-10 h. In such cases, however, the option exists of reducing the time by programming either the field or the flow. In the first case the spin rate is gradually lowered and the elution of late peaks thus hastened (Yang et al., 1974) and in the second case successive increases in flow rate at constant r.p.m. yield a rapid separation with little cost in resolution (Giddings et al., 1979). 2. Convenience of manipulation. Sedimentation FFF is a continuous flow separation technique and as such requires little attention once relaxation is completed and the eluant set in steady motion. Density gradient sedimentation on the other hand requires attention in the formation of the gradient and the loading of the sample. Once the sedimentation is terminated, particular care has to be given to the elution step in order to maintain the resolution generated by the centrifugation procedure. Elimination of the gradient former is a step requiring additional attention. 3. Amount of material processed. Sedimentation FFF is by its nature an analytical technique. At most only around 1 mg of sample can be processed in a run without encountering overloading effects and resolution loss. Larger samples require some combination of repetitive runs, parallel runs, wider channels, and some tolerance for resolution loss. For harvesting large amounts of material the density gradient sedimentation technique is advantageous since it lends itself readily to scale-up by merely increasing the radius of the gradient containing centrifuge tube. 4. Characterization of processed material. Density gradient sedimentation when pursued to equilibrium yields information about the density of separated components since these band where they are neutrally buoyant. By determining the density of the gradient solution at the position of banding a value for the density of the banded component is found for this particular chemical environment, that is, under the prevailing conditions of osmotic pressure and solvation, etc. Sedimentation FFF owes its selectivity to differences in effective mass. In those cases where solvent and sample densities are known with good accuracy this effective mass can be converted directly into molecular weight, or particle dimensions if the shape is known. Work with polystyrene latex beads has demonstrated a high degree of accuracy with regard to particle dimensions (Giddings et al., 1978). We now turn our attention to molecular weight values found in the course of this work. Our study of the gypsy moth nuclear polyhedrosis virus has involved 33 fractionations, which covers samples from over a dozen preparations. Effective molecular weight data obtained at a given time on a given preparation have been verified on several occasions by runs in a different centrifugal system whose channel thickness w is half as large (127 pm) as in the instrument described above. However, the calculated molecular weight for the first major retained peak in these fractograms shows a clear tendency to decrease

255

with time. Whether this is due to the presence of proteolytic or merely to the harshness

of the alkali-liberation

activity (Harrap et al., 1977)

process is not clear. In a few cases,

the effective mass of the first major retained peak (confirmed

by electron microscopy

to

contain rod-like structures) is low enough to correspond to envelope-free nucleocapsids. These values cluster around 78.3 X lo6 g/mol as recorded for the detergent treated rods. Leaving aside the runs yielding these low molecular

weights, as well as a small number of

runs showing substantially larger masses, we are left with 20 fractograms recorded under varying field and flow conditions which taken together give an average effective molecular weight of (160.6 f 16.0) X 106. The standard deviation can be considerably reduced if the analysis is limited to data collected on fresh preparations using a spin rate of about 1400 r.p.m. and a flow of 24 ml/h. An average of six observations collected under these conditions yields (160.0 + 6.8) X lo6 as the effective molecular weight. There is a slight apparent influence of field and flow conditions on the value obtained, but this effect is not well understood

since similar observations

on polystyrene

latex beads show very little

variation. By using the average effective molecular weight value obtained by sedimentation FFF, 160.0 X lo6 g/mol, in conjunction with the density value 1.192 g/ml obtained through isopycnic sedimentation, an actual molecular weight of 968 X lo6 g/mol is found. This value compares reasonably well with the value 900 X lo6 obtained by electron microscopic: studies (Bahr et al., 1976). It is expected that future sedimentation FFF work will eliminate the need for a density value from outside sources as runs can be made on one and the same sample in elution media of at least two different densities. ACKNOWLEDGEMENT

The authors wish to express their deep appreciation to Professor Hans Rilling for inviting our extensive use of his laboratory facilities at the Department of Biochemistry, University

of Utah. We are likewise indebted

of the Department

of Microbiology

This investigation

was supported

the National Institute

to Professor Don Summers and Dr. Don Witt

for helpful discussions. by Public Health Service Grant GM 10851-22

from

of General Medical Sciences.

REFERENCES Bahr, G.F., W. F. Engler and H.M. Mazzone,

1976, Q. Rev. Biophys.

Breillatt,

ME. Martignoni,

J.P., J.N. Brantley,

Appl. Microbial. Freifelder,

H.M.Mazzone,

9,459.

J.E. Franklin

and N.G. Anderson,

1972,

23,923.

D., 1970, J. Mol. Biol. 54,567.

Giddings,

J.C., 1976, J. Chromatogr.

Giddings,

J.C., F.J.F.

Yang and M.N. Myers,

1974,

Giddings,

J.C., F.J.F.

Yang and M.N. Myers,

1975, Sep. Sci. 10, 133.

Giddings,

J.C., M.N. Myers and J.F. Moellmer,

1978, J. Chromatogr.

Giddings,

J.C., K.D. Caldwell,

T.J. Dickinson,

Chem. 51.30.

125,3.

J.F. Moellmer,

Anal. Chem. 46, 1917. 149,501.

M.N. Myers and M. Martin,

1979, Anal.

Giddings,

J.C., M.N. Myers,

ed. D. Glick (Jown

K.D. CaldwelI

and S.R. Fisher,

Wiley & Sons, New York)

Harrap,

K.A., CC. Payne and J.S. Robertson,

Ignoffo,

C.M., 1968,

Bull. Entomol.

R., 1967,

in: Methods

of Biochemical Analysis,

1977, Virology

79, 14.

Sot. Am. 14, 265.

Lee, H.H. and L.K. Miller, 1979, J. Virol. Markham,

1980, in: Methods

Vol. 26, p. 79.

31, 240.

in Virology,

Vol. 2, eds. K. Maramorosch

and H. Koprowski

(Aca-

demic Press, New York) p. 3. Mazzone,

H.M., J.P. Breillatt

Pathology, Mazzone,

College

H.M.,

Invertebrate

and N.G. Anderson,

G. Wray,

W.F. Engler

Tissue Culture,

and GF.

Rigi-Kaltbad,

Summers,

MD.

Summers,

M.D. and L.E. Volkman,

and GE.

Smith,

1978, Virology, 1976, J. Virol.

M.N. Myers and J.C. Giddings,

Bahr,

Switzerland,

Ryel, E.M. and G.B. Cline, 1971, in: Methodology Reid (Longman, New Yrok) p. 149.

Yang, F.J.F.,

1970, in: Proc. IVth International

CoIloq.

on Insect

Park, MD, p. 371. 1980,

in: Vth

International

Conference

on

in press.

and Development

in Biochemistry,

84,390. 17,962.

1974, Anal. Chem. 46, 1924,

Vol. 3, ed. E.