Hiya rojer.  Good question.  The answer is an unequivocal:  yes and
no.  ;-)

Mostly, the virus isn't that quick, but you just never know when
it's gonna get up and go get quick.

Yes, if you want to reduce the risk of passing on an infection to a
partner, one of the ways to do it is take a hot, soapy shower just
before having sex.  However, like everything else that reduces the
risk of spreading herpes, there is still some risk.   And especially
while an outbreak is under way.  During outbreaks, yes, it is very
likely that the virus is pretty much continuously active.  During
other times, it's just a lottery, with your odds at around one in
twenty (though estimates there vary, too.)

And, of course, people don't have sex for only thirty seconds,
either, or at least I hope they don't.  If I recall, when a person
is newly infected, the virus travels from the skin, up the
nerveways, to the ganglia, at a speed close to five miles per hour!
That's fast, or so it seems to me, especially for a submicroscopic
ball of protein and no Nikes.  I have to assume that the little
buggers can wake up, travel the several inches to the surface cells,
and start replicating at about the same speed.  So, if you're going
to make love for half an hour, you need to be at least two and a
half miles away from your partner!

It works for me.

Take care,

Mike

Dude, you could even be shedding while your not having an outbreak. Just
assume your putting your partner at risk every time you have contact. The
risk may be very small... but I'm getting the feeling you're looking for a
loophole.

It depends of how recent you've had an OB.

A shower will definitelly help, but I cannot
tell if it will fully secure the interruption of the viral
transmission
even for the time immediately after having a shower if having visible
sores.
Try using topical Benzalkonium Chloride solution on top of the sores (
using a cotton swab); it seems
to penetrate deep into the skin-mucosa, killing viruses in deeper
layers.

These are issues, together with the shedding stuff, that are
controversial
in my views.
I do not understand the whole proccess of transmission, so far.

There is more information, if you check the site below,
with photos, graphics and additonal links to related articles in here:
http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Link&dbFrom=PubMed&f
rom_uid=11248101

(if you see below, on the left, it says (and links to related
articles, show:)

Here is the article:
http://www.pubmedcentral.nih.gov/articlerender.fcgi?tool=pubmed&pubmedid=11248101

I've also copy/pasted parts of it; from the lack of time
I cannot do a fair analysis right now, so to be
presented in the layman's terms for you.

However the article seems pretty down to Earth.

Perl Molson

Herpesviruses use bidirectional fast-axonal transport to spread in
sensory neurons

Abstract  
 
Alpha herpesviruses infect the vertebrate nervous system resulting in
either mild recurrent lesions in mucosal epithelia or fatal
encephalitis. Movement of virions within the nervous system is a
critical factor in the outcome of infection; however, the dynamics of
individual virion transport have never been assessed. Here we
visualized and tracked individual viral capsids as they moved in axons
away from infected neuronal cell bodies in culture. The observed
movement was compatible with fast axonal flow mediated by multiple
microtubule motors. Capsids accumulated at axon terminals, suggesting
that spread from infected neurons required cell contact.


Introduction  
 
Many viruses spread by infecting nerve terminals and traveling from
neuron to neuron in the vertebrate nervous system. The diseases
resulting from these infections are usually debilitating and often
fatal (1). Neurotropic herpesviruses (alpha herpesviruses) are an
exception in that typical infections are confined to the peripheral
nervous system (PNS) and are largely asymptomatic. Although the
herpesvirus genome remains in the PNS for the life of the host, and
newly reactivated virus made in neurons can retrace the path of
primary infection and return to the surface to be shed, alpha
herpesviruses rarely spread from the PNS into the central nervous
system. In instances when this does occur, the resulting encephalitis
has severe consequences (2). The mechanism by which alpha herpesvirus
restrict their spread to the PNS is not known. In fact, because
viruses are too small to visualize in living cells, the dynamics of
viral transport has never been directly assessed for any neurotropic
virus. Here we show that newly replicated individual herpesvirus
capsids (125 nm diameter) bearing multiple copies of the green
fluorescent protein (GFP) can be visualized and tracked in infected
neurons by laser-scanning confocal time-lapse microscopy. We
demonstrate that viral transport in axons is highly processive
(continues over large distances without stopping) and bidirectional.
Capsids travel long distances to ultimately accumulate at axon
terminals, suggesting that virions do not bud out of axon terminals by
a cell autonomous mechanism.


Materials and Methods  
 
GFP-Capsid Virus [Pseudorabies Virus (PRV)-GS443] Construction. The
PRV-Becker UL35 gene was cloned as a ≈4.5 kb SalI fragment (3),
along with the upstream UL34 gene, and both strands containing the two
open-reading frames were sequenced (GenBank accession no. AF301599).
The gfp open-reading frame was inserted into the PRV UL35 gene between
codons two and three, following a strategy previously used with herpes
simplex virus (HSV) type 1 (4). A recombinant virus, PRV-GS443,
carrying the fusion allele in place of the wild-type UL35 gene, was
made by using the pBecker3 infectious Escherichia coli clone (5).

Neuron Culture and Infection. Dissociated sensory neurons from the
dorsal root ganglia (DRG) of embryonic day 8–10 chick embryos were
seeded on 22-mm square glass coverslips pretreated with polyornithine
at ≈100 neurons/coverslip. The neurons were cultured for 3–5
days to allow for axon outgrowth before infection with the GFP-capsid
virus (6). Peripheral sensory neurons do not have dendrites in situ
(7), and only axons extend from DRG neurons in culture (8, 9). Axons
were identified as long projections that did not taper. Because the
number of neurons on a single glass coverslip was kept low to prevent
axons from contacting each other, much of the coverslip was exposed to
the viral inoculum. We found that the majority of the input virions
bound directly to the coverslip, as assessed by GFP emissions, and
never came into contact with the cultured neurons. To infect all
neurons on a single coverslip, each sample was incubated with ≈1
× 105 plaque-forming units of GFP-capsid virus for 1 h (incubating
with fewer plaque-forming units resulted in less than 100% of neurons
becoming infected; data not shown). Unbound virus was then washed away
before incubation was allowed to continue.

Confocal Microscopy. For time-lapse recording of living cells,
individual coverslips of infected neurons were sealed onto a glass
slide in Hepes-buffered media (pH 7.4) by using a 1:1:1 mixture of
Vaseline, beeswax, and lanolin. GFP emissions from infected neurons
were then imaged at 37°C with a Zeiss 510 laser-scanning confocal
microscope fitted with a heated stage and a heated 63 × 1.4 n.a. oil
objective. Excitation was at 488 nm with an argon laser, and up to
1,000 frames were captured per recording. For immunofluorescence,
coverslips of infected neurons were fixed, permeabilized, and reacted
with a mouse monoclonal anti-gB ("M2") antibody (10). The secondary
antibody was a goat anti-mouse conjugated to Alexa 546 (Molecular
Probes). GFP capsids were excited with a 488-nm argon laser, and Alexa
546 was excited with a 543-nm HeNe laser.

Quantitation of GFP Fluorescence. Individual GFP punctae observed in
samples of isolated capsids were measured for total emission intensity
by summing the values of all its pixels after first correcting pixel
values by subtracting background emission. The GFP emissions of
punctae seen in the axons of living neurons were measured in the same
way as above, except that the pixel values were corrected by
subtracting the emission background of the axon, which developed
notable background fluorescence during infection

Results  
 
A mature herpes virion comprises a DNA genome surrounded by an
icosahedral capsid shell made up of four virally encoded proteins. An
assortment of additional viral proteins, collectively called the
tegument, surround the capsid, which in turn is enclosed by a lipid
bilayer. Herpesvirus assembly and transport out of infected cells are
coupled, and only the capsid proteins are known to be continually
associated with the viral genome during intracellular transport. By
fusing GFP to a capsid protein, GFP signal remains associated with the
DNA genome of the virus during assembly and exit from infected cells.

We first cloned and sequenced the UL35 gene from PRV (GenBank
accession no. AF301599). This gene is homologous to the HSV UL35 gene
that encodes the VP26 capsid protein. Fusion of GFP to the N terminus
of HSV and PRV VP26 does not inhibit VP26 assembly into capsids or
substantially affect viral growth and spread in culture [(4); data not
shown]. Additionally, because there are 900 copies of VP26 per capsid,
individual GFP capsids produced sufficient fluorescence emissions to
be imaged with the short exposure times necessary to perform rapid
time-lapse microscopy.

Neurons were seeded at low density to prevent axons from fasiculating,
and only isolated neurons were examined. Axons of infected neurons
frequently contained many punctae of GFP emission. By using time-lapse
confocal recordings, some of these punctae were observed to move
directionally within axons (Fig. 1; see Movies 1–3, which are
published as supplemental data on the PNAS web site, www.pnas.org).
Moving punctae in axons were most readily detected between 8–14 h
postinfection (h.p.i.).

Although exact size determination was not possible, the observed
punctae were small and uniform in size, consistent with individual
fluorescent capsids. Although capsids are smaller than the spatial
resolution of light optics, they should appear larger than actual size
because of diffractive ballooning of the GFP emissions. This deduction
was confirmed by isolating capsids from infected pig kidney epithelial
(PK15) cells and dispersing the particles by sonication. The capsids
were then examined by negative-stain electron microscopy and confocal
microscopy (Fig. 2). By electron microscopy, 77% (n = 189) of all
capsids in the preparation were observed to be both intact and
isolated from other capsids by at least one capsid diameter (5% were
not intact, and 18% were in clusters of two or more capsids). We used
the same settings and optics for confocal scanning of the capsid
sample as used for examining living neurons. In this way, the GFP
emissions from 200 punctae were measured and plotted as a histogram
(Fig. 2). By assigning the lower 5% to emissions from capsid fragments
and the upper 18% to capsid clusters, we estimated an expected range
of GFP emissions for single isolated capsid particles. Comparison of
this emission profile to emissions from punctae in infected axons
indicated that the source of the punctae were individual viral
capsids. Although we occasionally observed motile punctae with
emissions 2- to 3-fold greater than expected for single capsids, large
clusters of capsids were never observed moving in axons.

Individual capsids were followed by using high-resolution particle
tracking to determine their location as a function of time. Successive
images of axons were captured at 0.1- to 1.0-s intervals, and
individual capsids were followed as they moved through the entire
field of view (typically 30 μm). Recordings were made with a bias
toward axons displaying significant capsid motility, and only axons
that were isolated from other neurons were analyzed. In this way, more
than 200 capsids were tracked from 28 time-lapse confocal recordings.
On average, capsids moved in an anterograde direction, away from the
neuronal cell body. However, capsids frequently reversed direction,
moving toward the neuronal cell body (retrograde travel) before
resuming anterograde travel (www.pnas.org). Some capsids moved
predominantly in the retrograde direction, but these also had periods
of anterograde travel. Because capsids could have a long period of
anterograde travel (>10 μm) followed by a long period of
retrograde travel, the population of retrograde moving capsids were
not necessarily distinct from the anterograde moving capsids. Capsids
moving in the retrograde direction were less common than those moving
in the anterograde direction, with a ratio of
anterograde-to-retrograde-moving capsids of approximately 7:1.

The processivity and velocity of capsid movement were quantitated by
examining individual capsid runs. A run was defined as a period of
uninterrupted travel lacking pauses or reversals in direction. To be
classified as a pause, motion needed to stop for more than a second.
By using these criteria, the average length of an anterograde run was
13.1 ± 0.6 μm (mean ± SEM, n = 198), and the standard deviation
of this mean was large (9.0 μm), as the distribution of runs
approximated a decaying exponential distribution (Fig. 3 A). This
number is an underestimate of the actual average travel distance,
because long runs were prematurely classified as ended when the capsid
moved out of the field of view (www.pnas.org). The average velocity of
all anterograde runs was 1.97 ± 0.06 μm/s (mean ± SEM, n = 198),
with top average speeds of ≈5 μm/s (instantaneous speeds
were sometimes greater) (Fig. 3 B). The average length of a retrograde
run was 6.8 ± 1.2 μm (mean ± SEM, n = 33), with an average
velocity of 1.28 +/− 0.12 μm/s. We note that capsid
velocities decreased slightly late in infection (>12 h.p.i.) (data not
shown).

Anterograde flux, or the frequency of capsid transport, was determined
by averaging the number of capsids entering the field of view (per
unit time) moving in an anterograde direction. The flux varied
significantly from axon to axon but did not change appreciably with
time postinfection. These observations may be explained by the
asynchronous infection of neurons within a sample and our bias to
collect data from neurons exhibiting significant capsid motility.
Bearing these considerations, the observed anterograde flux within
individual axons displayed a roughly Gaussian distribution (data not
shown) and ranged from 0.010 to 0.067 capsids/s, with a mean of 0.047
± 0.005 capsids/s (i.e., on average 1 capsid entered the field every
21 s). The average retrograde flux was 0.0072 ± 0.0015 capsids/s
(i.e., 1 capsid every 139 s). The distribution of capsids entering the
field of view was not random (i.e., χ2 analysis confirmed it was
not well modeled by a Poisson process), but rather individual capsids
had a tendency to travel in closely spaced clusters (data not shown).

Some capsids were observed to stall during axonal transport and never
regained motility during the remainder of a recording. In addition,
axons frequently exhibited stalled capsids that never moved during the
duration of a recording. The average number of stalled capsids per
micrometer of axon increased by roughly 0.16 capsids/μm/h.p.i.
(Fig. 3 C). Therefore, over a 30-μm stretch of axon, 4.8
capsids/hr were lost from the flow because of stalling. This
represents 3% of the total flow of 171 capsids/hr (based on the
anterograde flux of 0.047 capsids/s). The majority of capsids
accumulated at axon terminals (Fig. 4). To a lesser extent,
accumulations were also observed at varicosities, mid-axon, and in the
initial segment of the axon adjacent to the neuronal cell body (data
not shown). Axon terminals accumulated capsids along with the viral
structural membrane proteins including gB, gC, gE, and gI (Fig. 4 and
data not shown).


These results indicate that individual alpha herpesvirus capsids are
transported in axons at rates within the range of fast axonal flow
during exit from a neuron (11). Capsids are not transported as large
aggregates. Because fast axonal flow depends on microtubule motors,
capsid transport is likely mediated by two or more plus-end
microtubule motors. Even at frame rates approaching 0.1 frames/s,
anterograde capsid runs were longer than can be accounted for by the
processivity of single kinesin in vitro (12). The anterograde
velocities were modeled by a Gaussian curve, indicating that motion is
driven by a single class of plus-end motors. If two different types of
motors were involved, each with its own preferred velocity, we would
expect a curve with either two peaks (if their preferred velocities
were far apart) or distorted (if their preferred velocities were not
well separated). Although the criteria used in defining runs made
possible short undetected pauses that would give inflated processivity
figures, such pauses were deemed unlikely because of the observation
of numerous runs of 15 μm or longer whose instantaneous velocity
was never less than 2.5 μm/s.

A minus-end motor must also be present to account for the observed
bidirectionality of the capsids. Because newly replicated capsids move
bidirectionally during egress, but with net anterograde travel, we
hypothesize that during entry into neurons, capsids transported to the
cell body from an axon terminal would use a similar mechanism that
favors retrograde travel (13). We are currently investigating viral
entry dynamics in this system.

On average, the rates of capsid transport reported here are
approximately 3-fold greater than had been previously estimated for
HSV by using a dual cell culture chamber system (14). In the earlier
study, anterograde transport of virus from neurons in a central
chamber to epithelial cells in an outer chamber was estimated to occur
at rates of 2–3 mm/h (0.56–0.83 μm/s), on the basis of the
presence of viral antigen in axons near the epithelial cells. The
difference in rates is likely attributed to the assays used by the two
studies. The rates presented here are a direct measure of individual
capsid transport; in contrast, measurements made in the dual chamber
model are compounded by the kinetics of viral replication and viral
transport to axons from the cell bodies within infected neurons. Thus,
transport rates measured in the dual chamber model are likely
underestimates of actual capsid velocities.

To our knowledge, this work is the first direct measure of viral
anterograde transport kinetics in infected neurons for any neurotropic
virus. Although several studies have estimated the rates of retrograde
axonal transport of neurotropic viruses (15), a recent report has
directly measured the rates of HSV transport in the retrograde
direction by using severed segments of giant axon from the squid,
Loligo paelei (13). Although the squid is not a host for HSV, the
results indicate that retrograde transport of a viral tegument protein
fused to GFP occurred at rates averaging 2.2 μm/s. This rate is
similar to the 1.97 μm/s average velocity of anterograde moving
capsids during egress but is faster than the 1.28 μm/s average
velocity for capsids moving in the retrograde direction during egress
reported here. The faster rate of retrograde travel seen in the squid
model may reflect a difference in entry and egress kinetics or
differences in transport of viral tegument proteins and capsids.
Alternatively, because the virus likely uses host machinery for fast
axonal transport, the differences in retrograde rates may reflect
differences in invertebrate and vertebrate cell biology.

We observed that capsid and viral membrane proteins accumulate at axon
terminals and are not released. Alpha herpesviruses are competent to
spread from axon terminals of cultured DRG neurons to epithelial cells
(14) and spread transsynaptically in the chicken embryo nervous system
(16). Therefore, virus release from terminals may require cell–cell
contact, and the nature of the innervated cell may be a cue for
regulated herpesvirus spread in the nervous system. Because the
cultured neurons used in this report lack natural target cells, the
axon terminals are presumably growth cones (6). By using PRV
infectious clone technology (5), we are now in a position to identify
viral genes necessary for anterograde and retrograde transport in
axons, as well as for virion assembly and release from axon terminals.