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Why are sponges sometimes not considered multicellular?
I read somewhere (I can't find where) that there is no scientific consensus whether sponges should be considered multicellular organisms.
It seems I don't understand where is the line between unicellular and multicellular life.
I am not able to find a more elaborate explanation of that doubt. What are the reasons for it?

Sponges are generally considered as colonial organisms because there is little cell specialization and little separation of function/role. All cells do pretty much the same thing; it looks more like a pile of individual cells than an actual multicellular organism. In reality it is a little bit in between.
In any case, what one wants to call multicellular or unicellular is a matter of definition and preferences. You cannot find the line between unicellular and multicellular because there is no such line that would not be very arbitrary and filled with special cases.
You can study a little more the physiology of sponges and then decide for yourself if it looks sufficiently like a multicellular organism or more like a colony of cells (a colonial organism).

A collection of cells can maintain their individuality, and  have almost nothing in common but their location. No doubt all multicellular organisms today are descended from such collections.
To start doing anything useful for survival, the cells of such a collection can begin to specialize: some for locomotion, some for digestion, some for protection, and so on. Specialized cells group into tissues, and then into organs.
Organisms with cells responsive to electrical stimuli, which occur only in the animalia, can develop specialized tissue for movement, leading to muscles. They can also develop sensory organs, because excitable tissue gives them a mechanism for transmitting signals, and acting in response to sensory inputs.
A collection of cells without any of these specializations may develop into a syncytium, a mass in which there are  many cells sharing a common cytoplasm, with many individual cell nuclei lying in it. This arrangement allows them to share nutrients without having any specialized tissues or organs for doing so. Although there are multiple nuclei, it can almost be said that they live in a single, very large cell. Not having specialized tissues, they don't display the characteristics of multicellular organisms, and not having excitable tissue, they don't develop nervous systems.
This is the strategy some sponges have adopted, and that's why such sponges are not regarded as multicellular organisms.
Not all sponges are quite as simple as this, and have some differentiation in cell function. Whether these form tissues or not is a matter of definition.
So some sponges are almost entirely syncytial, and can't be regarded as multicellular, although they are metazoan. Some are a little more complex. And that explains why sponges will sometimes be regarded as multicellular, sometimes not. I guess it depends on the sponge.


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Why does my corn appear to have perfect flowers?
Corn is said to be a synchronously monoecious plant with separate male and female flowers.
I am growing a cross between a Peruvian and heirloom variety which seems to be growing some perfect flowers on both main stems, and suckers. I have googled this and find nothing to suggest this happens.
Is this common? And did a corn ancestor also have perfect flowers?

source: Stack Exchange

license: Creative Commons - Attribution Share-Alike - https://creativecommons.org/licenses/by-sa/3.0/

sort: votes


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aluminum sulfate vs (NH4)2SO4 in Colloidal Coomassie Staining Sol
I have been interested in Colloidal Coomassie Staining to detect proteins in PAGE gel.
I found 2 different recipes: one of them uses aluminium sulfate, the other, (NH4)2SO4. I am wondering about sensitivities of these recipes, and other advantages/disadvantages compering these two.
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3149902/
This article describes a recipe using aluminum sulfate.
http://onlinelibrary.wiley.com/doi/10.1002/elps.200305844/epdf
(NH4)2SO4 is used in this article.

all_licenses: Creative Commons - Attribution Share-Alike - https://creativecommons.org/licenses/by-sa/3.0/


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The largest Triceratops specimen
Almost every dinosaur book writes that Triceratops maximal length was about 9 meters (29.5-30 ft), see for example Thomas R. Holtz's dinosaur genus list 2011. My question is what specimen is the base or indication for that estimate, and what is its size? It could be a partial skeleton or maybe just a gigantic skull, and not a full vertebral-column specimen.
Also, what is the largest complete (or almost complete) skeleton of Triceratops? Peter Dodson in "The Horned dinosaurs" (1996) notes a 7.9 meters long skeleton at Minnesota's Nature Museum, but since then new almost complete skeletons have been found. If it is on display, please write in which museum it can be seen.
Added in edit:
The following image of EoFauna on DeviantArt and the discussion that follows suggests that Triceratops was as big as Eotriceratops, reaching a length of 9 meters and a mass of about 10 tonnes (10,000 kg) with skull about 2.7 meters long.
Added in edit 2021 December:
A specimen named "Big John" is 8 meters long and according to reports it is the largest mounted skeleton of Triceratops. It has 262 cm long skull and 1 meter long brow horns.

There's some references here https://en.wikipedia.org/wiki/Torosaurus. quote:  in 2006 Andrew Farke had pointed out that the new skulls described by him were on average even longer than Hatcher's original two: MOR 1122 has a length of 252 centimetres and MOR 981 of 277 centimetres.

here's a pic https://pbs.twimg.com/media/D6PV2vMVUAAVAmR.jpg

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Which extant species has the largest horns?
Among the bovids (cattle, antelope and all member of the Bovidae family): which species has the largest and longest horns? Please answer when including and excluding domesticated livestock and cattle.
I know that the Addax has about 85-110 cm long horns and they seem pretty long, but not as robust as a bull's horns.
Wikipedia writes that Greater Kudu has 120 cm long horns with a record of 187 cm (this figure, however, has a "citation needed" warning).
The extinct bovid Pelorovis had 1 meter horn cores and covered with keratin their length is estimated at 200 cm.
I saw in photos several (live) specimen of specially breaded cattle that had exceptionally big horns but I could not measure them.
For comparison, the longest Triceratops (an horned dinosaur) horns I know of are "Yoshi's Trike" 115 cm long horn cores with estimated total length of 130-150 cm when covered in keratin. However, the horns of Triceratops were thicker and more robust than any extant bovid's horns.

Why do you want to know something like this?

General interest. I am interested in numbers, records and records breakers.

Seems to me more Guinness Book of Records than biology, then, but who am I to judge.

all_licenses: Creative Commons - Attribution Share-Alike - https://creativecommons.org/licenses/by-sa/4.0/


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UACC-462 cell culture
UACC-462 is a pancreatic cell line established in the University of Arizona Cancer Center (UACC).
The cells are available from ATCC and there is the instruction for culturing UACC-462.
M-41 medium. The base medium for this cell line is ATCC-formulated Leibovitz's
L-15 Medium, Catalog No. 30-2008. To make the complete growth medium, add the following components to the base medium (final conc.):
fetal bovine serum to a final concentration of 5%
0.01 mg/ml transferrin
0.01 mg/ml insulin
5 µg/ml (55 U/ml) catalase
3.6 µg/ml (0.01 mM) hydrocortisone
10 ng/ml EGF
3 ng/ml Estradiol
70 µg/ml O-phosphorylethanilamine
0.0008 µg/ml L-thyroxine
extra 2mM L-glutamine
Atmosphere: air, 95%; carbon dioxide (CO2), 5%.
Temperature: 37°C
On the other hand, L-15 is designed to culture without CO2 and often used time-laps experiments under microscope.
My question is why L-15 based media can be used in CO2 incubator. Is the cell line acid-philic?

There are a number of cancer cell lines that are grown in various forms of L-15 in 5% $CO_2$ incubators. I don't know the exact biochemistry behind it, but it works, so that's what's used.

@MattDMo Regarding this cell line, an ATCC technician admitted the mistake. Do you have experiences culturing cancer cells with L-15 based media in 5% CO2? I would like to know which cells can be grown under the condition, for future reference.

unfortunately, I am no longer at the lab where they did this, so I can't check the database. So, ATCC said the line shouldn't be cultured with $CO_2$?

@MattDMo "It should read 100% atmospheric conditions." That's what they said.

@MattDMo May I post another question to ask cells that can be grown with L-15 based media in 5%CO2?

I suppose you could, but it seems like a fairly narrow question. I'd do some good googling for a while to learn the history of L-15 and see if you can find some lists on your own. A lot of labs post their protocols online, so you may find something there.


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PAGE gel in EtBr
I have been using TBE-PAGE gel to see small fragments of DNA by post-staining that means gel is stained with EtBr after electrophoresis.
Someone told me that you can add EtBr before polymerizing acrylamide, but I have never seen protocols like that.
Have you added EtBr before polymerization? Have you compare these staining methods? Let me know your experiences and knowledge.

TBE-PAGE gels generally take a while to run (especially the big Urea-PAGE gels for RNA) and the EtBr can quickly migrate out of the gel. Since PAGE gels are relatively thinner than agarose gels, staining post-run is not a problem. However, I have not really tried adding EtBr before polymerization.

You can also add some EtBr to the running buffer in the anode chamber. The EtBr is positively charged and will migrate through the gel towards the cathode while the nucleic acids in your sample will migrate towards the anode.

Thank you for the comments, all. I seems you can add EtBr in gel before polymerization. And I could see the problem when doing so, and the solution.


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Were there any marine/aquatic dinosaurs?
Most dinosaurs were terrestrial, but there were a couple of groups of arboreal and flying dinosaurs (microraptors, birds etc).
I have read that the theory that Brachiosaurids were aquatic has been largely discredited1, but that Spinosaurids were probably semi-aquatic2.
Excluding modern marine birds (e.g. penguins etc.), were there any other marine/aquatic dinosaurs?

Note: I am aware that the marine reptiles mesosaurs, phytosaurs, mosasaurs, dolichosaurs, Icthyosaurs, thalattosaurs, Sauropterygia (placodonts, nososaurs, plesiosaurs etc), Choristodera were not part of Dinosauria.

References:

Three new sauropod dinosaurs from the Upper Jurassic of Colorado, Great Basin Naturalist.
The Physiology of Dinosaurs: Circulatory and Respiratory Function in the Largest Animals Ever to Walk the Earth, Respiratory Care
A note on the habits of sauropods, Annals and Magazine of Natural History
Tipsy punters: sauropod dinosaur pneumaticity, buoyancy and aquatic habits, Proceedings of the Royal Society of London

Oxygen isotope evidence for semi-aquatic habits among spinosaurid theropods, Geology
Semiaquatic adaptations in a giant predatory dinosaur, Science
Convergent evolution of jaws between spinosaurid dinosaurs and pike conger eels, Acta Palaeontologica Polonica
Functional Morphology of Spinosaur 'crocodile-Mimic' Dinosaurs, Journal of Vertebrate Paleontology

How about the mosasaurs for example?

@Remi.b mosasaurs were squamates, not dinosaurs: Mosasauroid phylogeny under multiple phylogenetic methods provides new insights on the evolution of aquatic adaptations in the group, PLOS ONE

Fair point! I am wondering why this post has been down voted.

Hesperornithes could be a good example but they are Aves (a "basal branch") if I am not mistaken. They are not modern birds though. Would you accept it as an answer?

@Remi.b ah, thanks! I was thinking of pre K-T extinction dinosaurs when I posted the question, which this is, so yes I would - unless someone responds with a larger list of examples :)

There are a couple of specimens of a dinosaur species named Liaoningosaurus paradoxus that have been found with fish in their stomachs and skeletal features suggesting it was at least semi-aquatic.  It is a member of the ankylosaurian dinosaurs from the early Cretaceous period of China.
From Fish Hunting Ankylosaurs( Dinosauria,Ornithischia) from the Cretaceous of China:

With elongate and fork-like denticles of cheek tooth crowns,L.
  paradoxus has a dentition capable of penetrating into animals like
  small fishes. The carnivorous adaptation of the dinosaur is also
  supported by the ungual modification to a sharp claw in both the
  fore-and hind-limbs. The evolution of a shield-like ventral armor
  plate and the loose sacrum-pelvic connection suggest that L. paradoxus
  may have adopted an aquatic way of life,using the ventral armor plate
  to protect the body from underwater attacks; as such,the open suture
  between the neural arch and centrum of the vertebrae cannot be used to
  indicate the juvenile nature of the type specimen. L. paradoxus is the
  first carnivorous ornithischian dinosaur since dinosaur was first
  known in the 18th century and represents not only the first aquatic or
  semi aquatic example of armored dinosaurs but also the smallest
  species of ornithischian dinosaur so far known.

Image Source: [Paleontology • 2016] Liaoningosaurus paradoxus • Fish Hunting Ankylosaurs (Dinosauria, Ornithischia) from the Cretaceous of China

I found this wikipedia page which lists a few more examples:

Only a few nonavian dinosaurs are thought to have been semiaquatic. The combination of being oviparous and endothermic seems to have prevented the evolution of fully aquatic dinosaurs.

Theropoda
Paraves

Avialae

†Ichthyornis - a toothed ornithuran analog of modern seabirds such as gulls and petrels
†Hesperornithes - a mostly flightless group of diving bird-like ornithurans

†Dromaeosauridae

†Halszkaraptor - a mallard-sized basal dromaeosaurid from Mongolia with flipper-like forelimbs

†Megalosauria

†Spinosauridae - thought to be piscivores, the group had crocodile-like skulls and includes some of the largest known carnivorous dinosaurs

†Ornithischia
†Ornithopoda

†Hadrosauriformes

†Lurdusaurus - an unusually heavy-bodied and short-limbed iguanodont conjectured to have been similar in lifestyle to the modern hippopotamus

List of semi-aquatic tetrapods: Non-avian dinosaurs

Chapter 23 of New Perspectives on Horned Dinosaurs: The Royal Tyrrell Museum Ceratopsian Symposium, "A Semi-Aquatic Life Habit for Psittacosaurus, describes the evidence suggesting Psittacosaurus was semi-aquatic.
P57-59 of Discovering Dinosaurs: Evolution, Extinction, and the Lessons of Prehistory suggests some hadrosaurs may have been semi-aquatic:

Dinosaur
Illustration

Icthyornis

Hesperornithes

Halszkaraptor

Spinosaurus

Lurdusaurus

Psittacosaurus

The only example I could find for the moment are Hesperornithes. Hesperornithes are actually part of the Aves clade (see here on tolweb.org) but they are not modern bird species as they existed during the Cretaceaus.

Note that there are examples of aquatic species in Squamata (Squamata are not Dinosauria) though such as the mosasaurs for example.

According to the latest discovery, Nature (Published: 29 April 2020), Tail-propelled aquatic locomotion in a theropod dinosaur, Spinosaurus was probably aquatic or at least semi-aquatic. See also this video.


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What is the minimum number of clades necessary to partition Vertebrates while maintaining the classical groups of Birds, Mammals, and Amphibians?
At school I was taught the Linnaean view of vertebrate life - that vertebrate animals consisted of 5 distinct groups: mammals, birds, reptiles, amphibians, fish.
I learned later however that this was phylogenetically inaccurate, with birds being 'closer related' to crocodiles than other reptiles are, for example.
As such I am curious - since the idea of these groupings is so strong in popular conceptions of animals:
What is the minimum number of clades we would need in order to partition 'Vertebrates', maintaining 'Mammals', 'Amphibians', and 'Birds' as clades?

@Remi.b By completely partition I mean, every member of vertebrates belongs to one of the clades, and the clades are disjoint; by minimum I mean make the remaining clades as large as possible given the constraint Birds, Mammals, Amphibians are already clades. e.g. Crocodilians is 1 clade (considering smaller clades e.g. Crocodiles, Alligators & Caimans just increases the amount we need to consider, therefore not minimum), and it cannot be extended to form a larger clade as this would be (at least) 'Archosaurs', and hence absorb 'Birds', breaking one of the conditions.

Oh I got it now $\ddot \smile$. Here is the beginning of the tree of vertebrates and as you can see there are already quite of few branches that will need named to "completely partition the clade of vertebrates". One could just screen through tolweb.org (or some other similar ressource) and count.

Phylogenetic taxonomy of the Vertebrates (mun.ca) has the following graphic (14 clades total, minimum of 12 collapsing Amphibia):

Here, what was preivously 'Fish' (the paraphyletic group of non-tetrapod vertebrates) becomes 6 clades, and 'Reptiles' (the paraphyletic  group of non-avian Sauropsids) becomes 3.
It seems the question is equivalent to asking "what is the minimum number of clades necessary to partition Vertebrates while maintaining 'birds' as a clade?".

Related: https://biology.stackexchange.com/questions/88511/is-a-lion-a-bony-fish


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Insect (I hope) identification
Look, let's get something straight.  I am not the worlds biggest sissy when it comes to bugs, but I'm not David Attenborough either.
Now that that's out of the way, down to business.  Living in Michigan as I do, I was quite surprised to find what looked like the spawn of Predator and a cicada living in my back yard.   Suffice to say it made me nervous.  In one of the pictures below, I have included a cigarette with about 3 drags smoked off of it for sizing.
Please offer insight into the true nature of this animal.

Looks like a crayfish to me. Did it smell bad? It looks pretty dead, was it in fact dead? Do you live near a river?

Yes.  Almost certainly dead.  Or very very chill.  I don't live near anything remotely resembling a river.  Where the picture was taken is in an electronics box just under a panel in my yard.  Not deep (10 inches tops).  Box is covered.  This is what I saw when I lifted it.

How far away is the local water table? if you live in a wettish area it's probably a prairie crayfish. They're local to the region, apparently really common, and will live anywhere the dirt is wet. Or damp. (also, have you ever seen a live lobster?)

I have seen a live lobster.  Not on land though.  I guess I live in a wettish area (it's on a golf course and there is varying terrain), but there is (too my knowledge) no immediate source of water within a couple 100 yards.  In retrospect, it does look like a crayfish; I just was a bit shocked to find one in what I thought of as a relatively dry area.

Now that I think about it, that may be why it's dead. They're great for the soil, at least.

This is not insect at any rate )))

In the absence of someone more qualified, I believe this to be a prairie crayfish, based on where it was found (land) and what it is (decapod of some kind that looks like a crayfish). Therefore it is unfortunately not an insect.
It's in the right range and roughly the right terrain.
I welcome entomologically qualified advice!

Yes, definitely a crayfish (Crustacea) and not an insect. Both groups belong to Arthropoda though.

Well; I guess it's okay that it's not an insect since it's a crayfish.  I was more concerned about it being one of Satans minions. : [


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Why does radiation sickness cause hair to fall out?
As I understand it, radiation poisoning (acute radiation syndrome) is fatal because the radiation kills cells. However, why do victims experience hair loss at such an early stage in the process?

Any link at hair loss? I doubt it. Hair loss may be due to chemoterapy if the affected persons contracted cancer or leukemia.

Radiation levels used for treatment of cancer does kill nearly all hair in the target zone.

Radiation poisoning causes mutations in DNA that affect normal cell function, often causing them to die.  Cells normally have a number of repair mechanisms but if the damage is too great they won't be able to do so.  In particular, cells that are dividing quickly will not have time to repair their DNA before division and so die far quicker than other cells.  Chemotherapy is built on this concept.  By selectively introducing drugs or radiation that target fast-dividing cells, the cancer cells, which are very dividing rapidly and unchecked, are killed off before most normal cells.
However, there are some cells in the body that are very rapidly dividing.  Hair cells are one such type, hence hair will often fall out during chemotherapy or radiation sickness.  Another big cell type are those in the gut, which are constantly dividing.  That's why you often see people in movies who have been exposed to radiation vomiting.  The radiation targets those cells first, hence those are the first symptoms.

I suppose that in any radiation exposure, the skin is what is damaged first. So radiation burn on your scalp, may be the primary cause for acute hair loss. Possibly because of the lower penetration depth of α-radiation, which is more damaging.


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Are there any documented instances of coronaviruses being directly transmitted from bats to humans?
Many human coronaviruses have ancestral host origins in species of bat. However, all instances I am aware of identified other animals as intermediary vectors:

SARS-CoV: Human ← Palm Civet / Raccoon ← Bat
MERS-CoV: Human ← Dromedary Camel ← Bat
HCoV-229E: Human ← Alpaca ← Bat

Are there any known cases of direct bat to human transmission of coronavirus?

Source: Bats and Coronaviruses, Viruses (2019)

Have you ever thought of "identified other animals as intermediary vector" as a kind of necessity? An  intermediary vector is a necessary, searched for missing link for the very fact that bat virus strains just do not infect human cells (recent research in Cambodia found bat virus strains with even better affinity to target receptor ACE2 - it is not at all clear  if those species were able to infect a human cell. "I guess not!")

It is possible, but unconfirmed, that the first incidents of SARS-CoV were directly infected by bats, and that palm civits were infected parallelly:

The new study demonstrates that Chinese horseshoe bats harbor a pair of SARS like viruses that are very closely related to the strain that caused the 2003-2004 pandemic and that these strains have the capacity to directly infect human cells and civet cat cells as well as bat cells. These findings suggest that bats may have directly infected humans and that civet cats may have been incidentally infected.

More Evidence Emerges That Bats May Have Spread SARS, JAMA (2013)


text:
How do bats survive their own coronaviruses?
How do bats survive their own coronaviruses (without showing any symptoms)?
Or, more generically, how can viruses keep reproducing inside healthy carriers without inducing any pathogenic effect?
Are coronaviruses able to replicate themselves without harming bats, or maybe these viruses are just latent like herpesviruses in humans?

Related question: Why are bats the source of dangerous coronavirus pandemics?

Perhaps bat immunity is a result of their crowded living conditions needed to survive cold winters.  The bat that gets sick and dies does not breed.

It's common for the reservoir host of a zoonotic virus to be tolerant of it. MERS coronavirus appears to cause mild or no disease in dromedary camels ( source ), but kills about 35% of confirmed infected humans. ( CDC ) Sin Nombre hantavirus seems to be mild in the deer mice that spread it, despite ~36% fatality rate in humans. ( source ) Mosquitoes are efficient vectors for flaviviruses like dengue and zika in part because they have adaptations we lack that protect them from the virus. ( source ) Also, human communities are host to several viruses: about 90% of people have a herpesvirus infection ( source ) with similar numbers for polyomaviruses. ( source ) Very few of these infected individuals show symptoms.
The wide prevalence of these asymptomatic infections shows that the virus is successful when it can replicate while the host remains healthy. In general, virus reproduction kills cells, and when cells die faster than the host can replace them, this causes symptoms up to and including death. The host's immune system supresses virus activity, and a virus that can't evolve ways to avoid immune supression will be wiped out. But if the virus gets so good at avoiding the immune system, it will kill the host, which is bad for the virus. So both host and virus tend to evolve to a point where the immune system wins most, but not all of the time. There is a lot more to it, but I'll stop here.
In conclusion, we don't really need to look for special properties of bats to explain their tolerance of coronaviruses, even though, as iayork points out, there are reasons to expect bats might be more resistant.

this is why I added these lines to my question: "Or, more generically, how can viruses keep reproducing inside healthy carriers without inducing any pathogenic effect?"

Excellent answer. As Lewis Thomas said: "Disease usually results from inconclusive negotiations for symbiosis"  The question is less one of why the bats don't get sick and more why we do.

@Zafalija I've added another paragraph which I hope answers this part of your question.

@timeskull I was rereading your answer: "There is a lot more to it, but I'll stop here" Could you link me something about virus-host evolution?

@Zafalija Here's the best review I could find: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4207818/

It's been tentatively proposed that bats are often unusually able to tolerate long-term infection with a wide range of viruses (though this hasn't been formally shown to be true). A specific cause for this (possible) phenomenon isn't known, but many possible explanations have been put forward. It's likely that if it's true, there's no single cause but rather a combination of multiple causes to add together.

Bats have an array of unique life history characteristics that not only allow them to be particularly good reservoirs for viruses that are highly pathogenic in other species, but also appear to have shaped their immune systems. Although research on bat antiviral immunity has focused on only a few species to date, at the genomic level, selection on genes is concentrated on the innate immune system across both suborders of bats. However, while these studies have provided a rich source of hypotheses, the majority remain to be tested at the functional level and many questions remain that cannot be answered from comparative genome studies. Experimental studies to date have demonstrated some functional differences between bat species, with the common emerging theme that the overall antiviral response appears to converge on a lower inflammatory profile, with tight regulation of the cytokine and inflammatory response key to clearing viral infection without the pathological outcomes typically associated with infection.

--Going to Bat(s) for Studies of Disease Tolerance


text:
What is the formula to calculate $R_0$ (basic reproduction number)?
What is the mathematical formula for $R_0$ and what does each variable represent?
I have tried searching this to no avail.

$$R_0 ∝ \left(\frac{\text{infection}}{\text{contact}}\right) · \left(\frac{\text{contact}}{\text{time}}\right) · \left(\frac{\text{time}}{\text{infection}}\right)$$
More specifically:
$$R_0 = τ · \bar{c} · d \quad \quad (1)$$
where $τ$ is the transmissibility (i.e., probability of infection given contact between a susceptible and infected individual), $\bar{c}$ is the average rate of contact between susceptible and infected individuals, and $d$ is the duration of infectiousness.

Notes On $R_0$ (2007)

Further reading:

https://en.wikipedia.org/wiki/Basic_reproduction_number
Complexity of the Basic Reproduction Number (R0), emerging Infectious Diseases (2019)
Theory versus Data: How to Calculate R0?, PLoS One (2007)
The R0 package: a toolbox to estimate reproduction numbers for epidemic outbreaks, BMC Medical Informatics and Decision Making (2012)


text:
Sequence analysis software suites
I have used DNAStar laser gene and VectorNTI in the past for cloning, primer design, sequence alignments, etc but no longer have access to these. Are there any comparable replacement suites or what stand alone packages would you now recommend?

TO do what? Please explain your actual requirements.

And for which operating system?

I should have specified OS. Either OSX or Linux. As for what I want to do with it, I need to be able to at least organize sequences, design primers from large sequences, local alignments, make plasmid maps, search ORFs, develop cloning strategies, etc. I also want to read chromatograms

All of this you can do with benchling (see answer below). And as it is web based it is not operating system dependent. As far as I know Geneious and GeneDesigner don't work in Linux.

CLC workbench is a decent program as well (see response below).

I would strongly recommend benchling https://benchling.com
This is an awesome web based tool for cloning, primer design, multiple sequence alignments and everything else you are used to doing with the other tools. It is very user friendly, and most importantly you can share designs with your collaborators. Also, the graphics are very beautiful. I recently started using it and I am really impressed with the technical support they offer: you can chat with them and they would reply within minutes. Also, if there is a feature you would like to have implemented they would definitely consider it. Benchling is free, open source and you might be even able to code up some features yourself.

My pleasure! Are you a benchling user?

Starting from today.

I personally use more frequently Geneious for most of the basic every-day manipulations (my university bought a license), but I would recommend Ugene: it's free, open-source, cross-platform and supports batching and scenarios.

I would recommend Galaxy. https://usegalaxy.org/
The system does include primer design, sequence alignments and covers many common bioinformatics tasks.

For primer design you can use the Gene Designer: https://www.dna20.com/resources/genedesigner
For sequence alignment you can use BLAST (from the NCBI, also downloadable as a standalone tool)
Of course there are better options that might be more specific for your situation.

If it's not loo late to contribute, CLC workbench is nice too (http://www.clcbio.com/products/clc-main-workbench/). I only use the free version (http://www.clcbio.com/products/clc-sequence-viewer/) but its very user friendly in terms of the features I have worked with.

Geneious is probably the strongest standalone package, but real bionformaticians rarely deal with GUI driven programs.

Why do you say that "real" bioinformaticians rarely use GUIs? Is it just some sort of macho "I'm too cool for that user-friendly stuff" attitude? REAL anythings get the job done with the best tools that are available for their particular problem domain. Sometimes it's text-based utilities, sometimes web-based, sometimes GUI-based. It's not about how much pain you can inflict on yourself, but whether you can get the job done correctly, on time, and on-budget.

@MattDMo It's not just elitism. For sequence manipulation, GUIs are much less efficient. They only make sense for people who have so little competence with programming that the added cost of learning this makes suffering a GUI desirable.

@Superbest please back up your statement with data that GUIs are "much less efficient." With today's modern multi-core computers and huge amounts of available RAM, the cost of running a graphical interface is not terribly high, considering that most users will be using one anyway. I am perfectly comfortable using Linux through the command line, and I am rather proficient in Python, but if pointing and clicking makes a one-off job easier than writing a new program for it, I'll do it. You say it's not elitism, but the rest of your comment is as elitist as they come.

They're not inefficient of resource usage, they are inefficient in terms of user time and effort. Many GUI tools are also critically underpowered. The Geneious next-gen alignment functions, for instance, do not output a SAM file (to my knowledge) that you can manipulate with other tools if Geneious functionality is not enough for you. Adding to that is the issue of citing proprietary algorithms in your publications.

@MattDMo Also, I'd like you to explain how my comment is "as elitist as they come".


text:
What is the name of this moth found in Virginia USA?

Found in Virginia, USA on Nov 1 2015, 10:34 AM.

Oh, found it. It is called a Euonymus Leaf Notcher Moth. It is an invasive species from Asia. It was first found in Maryland and Virginia in 2001.
http://www.insectid.ento.vt.edu/exotic-pests/euonymus/


text:
Is this an instar of the red fire bug?
While inspecting some yarrow that seemed to be having problems, I found a few of these bugs. I think they may be an instar of red firebug, but the photos of the nymphs that I found had only one row of black dots (e.g. here along the back) whereas these have two, maybe more. (US, midatalntic, tiny!!! Maybe 1.5 mm?)
Are these firebugs or something else? (Sorry for the poor quality photo.)


text:
mouse gene translated to human gene
I have a list of mouse genes, but all our analysis happens for human genes. Is that possible to translate these mouse genes to human genes? and is there any tool that can help me on this?
I am mostly a computer science guy, I am not sure this is even possible. However, according to someone, this could happen, I don't expect a full conversion though.
Thanks.

Your question is a little unclear. Are you trying to find the sequences for the corresponding human genes, or just the name of the human gene?

I would recommend using Biomart from Ensembl. This is basically a Swiss Army Knife for converting gene names into various IDs, getting corresponding locations on the chromosomes and so on. You can upload a list of gene names and convert them into the corresponding gene IDs of other species and the convert these IDs back to gene names.
If you haven't used Biomart yet, you can either start by going on their help page or by having a look on this tutorial for the conversion of IDs.

Yup, use biomart.  When you say you have a list, do you have a list of ensembl gene IDs, or a list of Gene Symbols, or Refseq IDs?  The gene symbol is probably what you want; the name that you will be able to find in both genomes  ("Cdh23" or "ApoE1" are examples).  But of course there will be some mouse genes that aren't in humans, and vice-versa.

Sounds like you are searching for homologs of mouse genes in human genome. BLAST should be able to help you finding that. You might want to find the sequences for the mouse genes on GenBank and run BLAST against it.
However, in this scenario, protein BLAST (BLASTp) will actually helps you more since the protein sequences are conserved better than DNA sequences are between species. In this case, you need to find the proteins coded by the mouse genes, and run BLASTp then.

Depending on how long the list is, you don't want to use Blast. Biomart (once you learned how to use it) is a really good tool here.


text:
How many photons does a GFP molecule absorb per second?
I'm trying to calculate how many photons will a GFP emit per second. The calculation seem easy just multiply the number of absorbed photons by the quantum yield. But how can I obtain the number of absorbed photons?
I know this would be directly related to the absorbance of the protein and the light intensity.
Is there an equation that relates both parameters to obtain the number of photons absorbed?
GFP absorbance at 488 nm:
$A_{488}=5.6\times10^4M^{-1}cm^{-1}$.

This paper is behind a paywall, but the supplementary material is available for free. In the last paragraph of page S-3 they seem to be calculating what you're after, referring to three different papers as their sources. Goes beyond my (bio-)physics knowledge to comment on the accuracy though.

Since the quantum yield isn't the main focus here, I'll just take the Wikipedia figure of 0.79 photons emitted / photons absorbed
The rest can be worked out from first principles.  The absorbance represents the number of log10 units by which light is reduced, but its units are designed to confuse:  5.6E4 (L/mol) / cm = 5.6E7 cm^3/mol / cm = 5.6E7 cm^2/mol GFP.*  This represents the area you can paint with 1 mol of GFP to reduce light passing through by 90%.  But let's substitute the value of a mole, to get 9.3E-17 cm^2 = 0.93 A^2 per GFP.  That chromophore casts a nice little shadow.
But our units aren't giving us photons - why not?  Well, the absorbance and fluorescence is proportional to the illumination.  So we need some notion of the amount of light, preferably something like irradiance (W/m^2).  The first figure I saw on a search was about 300 kW/m^2; you'll need to decide on your own.  So take 9.3E-21 m^2 * 300 kW/m^2 = 2.8E-15 W absorbed per GFP assuming this 90% absorption level.**
Now we still have to figure out the joules per photon.  The frequency of this light is (3.0E8 m/s)/488 nm = 6.1E14 / s = 610 THz (hmm, maybe a little more cyan than green).  We take that times Planck's constant = 6.63E-34 Js and get 4E-19 J/photon.  Divide 2.8E-15 (J/s)/GFP by that and we have 7000 photons per second passing through GFP, of which we're supposing 90% are absorbed.   And then I have to also multiply by 0.79  quantum efficiency.  Taking those together it is approximately 5000 photons
I encourage you to rework this since there's some chance of foul-ups here, and remember to get your own numbers for the microscope irradiance.
*Remember absorbance is -log10(transmittance), so an absorption of 1 means 90% of the light is blocked.  So the 56,000 there indicates that 1 mole of GFP can cut the light by 90% ... 56,000 times over again.  Or we can divvy it up into 56,000 different square centimeters and reduce light in all of them by 90%.
** 90% of this will be absorbed at a painted surface, such as a slide.  In a cuvette the energy absorbed would be much less because most of the GFP is in shadow, while spreading the GFP out further would increase the photons absorbed a bit because the rear part of the 90% is still in shadow.  I think 90% absorption isn't tremendously unreasonable to use since people will try to arrange for as bright a signal as they reasonably can, which may be localized to small features on a slide; but for example absorbing 68% of photons (reducing transmittance by the square root of 10 with an absorbance of 0.5) would cover twice the area, collecting 36% more photons total.  The limit at 0% would be ln(10) = 2.3 times higher than the figure at 90% absorption, but you can't see GFP if it's spread out to 0% absorption.

Hi @Mike_Serfas I think you nailed it. Just as a favor to people that will read this in the future could you please explain in more detail the way you get the per molecule absorption (specially how you go from absorbance to area). Also could you change the format so the equations are displayed in a diferent row?

@BPinto - thanks for making me look at this again.  Spreading out the GFP more varies the amount of light absorbed by more than I was thinking, though I should have known, because at 90% much of it is in shadow.  I'm not sure it's wrong to assume 90%, but there's no two sigfigs to be had here.  As for the equations ... well, I don't enjoy trying to code those formats when the text-based version seems so practical.


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What options are available for high-throughput isolation and identification of microbes from soil?
I have a lot of experience with isolating individual colonies / species / organisms from the environment and then sequencing them for identification, but the process is slow and cumbersome.
My goal is to build an organism library so actually having the individual isolates is desired. The metagenomic approaches are great for screening a pool of organisms for something I'm looking for, but I still have to separate it out if I want to keep it.
I currently create soil dilutions, plate them, isolate colonies early on to create axenic cultures and then DNA barcode the microbes. This process is time-consuming and often biased towards faster growing microbes.
Are there any high-throughput methods of isolating and identifying microbes (mainly fungi and bacilli) from a mixed-substrate like soil?
A literature search on this subject doesn't reveal many options that are currently accessible. I'm seeing some research on microfluidic devices that may help, but I'm not aware of any microfluidic platforms that are available to consumers.
Aside from this technique and microfluidic techniques (not currently available to my knowledge), are there any other techniques to consider to improve throughput?

Can you please provide more detail and focus your question a bit more narrowly as this kind of open ended question isn't a good fit for this site. (e.g. Do you really need to isolate and why?) ——— If I search for "soil genomics" the top two hits are articles from Nature Reviews Microbiology titled "Soil genomics" and "The metagenomics of soil". Have you read these and looked for more recent reviews and articles that reference them? Does this not allow you to find answers?

The question is very specific: what high-throughput methods are available for isolating and identifying microbes like fungi and bacteria? I don't know how to make it more specific than that. I stated what I want to do which is separate (isolate) the microbes and identify them (their species). I would like to know what high-throughput methods are available. The reason why has nothing to do with the question.

Have you looked into metagenome-assembled genomes? E.g. don't bother with isolation, just process everything together and rely on data analysis to pull the different genomes apart. This can be done with or without extra lab steps that add additional info (I once worked at a company that did this as a service: https://phasegenomics.com/products/proximeta/, so conflict of interest there). This is pretty routine at this point. You can also do it on standard shotgun metagenomes (e.g. https://www.nature.com/articles/s41587-020-00777-4), it's just slightly messier.

I'm building an organism library so actually having the individual isolates is desired. The metagenomic approaches are great for screening a pool of organisms for something I'm looking for, but I still have to separate it out if I want to keep it.

Do you now see why I suggested you tell us why you want to do this? SE sites get many questions that fall into a trap known as the X-Y problem, which means it is almost always a good idea to include "why" information in your questions. It is also best to include such information in the text rather than responding in comments — this is because comments are ephemeral and often overlooked. Finally edits should be incorporated directly into the text and not labeled — changes are tracked by the site anyways.

If your goal is just to get a lot of unique isolates, picking colonies from agar plates is pretty much a necessary evil. Colony picker robots can automate that part of the process. One potentially faster alternative to DNA barcoding is whole cell MALDI-TOF-MS, which can dereplicate isolates and potentially ID some of them with medium-ish throughput. I can pick and screen about 1000 colonies in a day without automation. Most of the ID libraries are geared for pathogen ID, so you'd probably end up sequencing still. But you can add your own spectra as your collection grows.

I've looked into MALDI-TOF but it seems like there are two big issues: one is that even if there is a strain of interest found in the results, I still need to go through the process of plating and isolating it and hoping I can distinguish it morphologically from the rest. The other is price. I'm an indie scientist and can't spend $20K on a machine like that. Curious to hear your thoughts on it though. Appreciate the info for sure.

@tyersome I can see your perspective clearly, yes. Can you understand how I might also see from my perspective that the question was clear? I don't think either side is all right or wrong. Clearly, SE admins/users/mods are trying their best to deal with an onslaught of low-quality questions. But as a long-time user, I am frequently met with downvotes and negative reactions for not asking a question in just the right way. It's a negative experience for both of us. I don't know the solution but I'm not alone in these thoughts.

@MikeyC I'd love to pick your brain more if you'd allow for it. Is there a way I can connect with you? I'd like to learn more about how you're able to screen 1000 colonies without automation. And what does screen mean exactly? (ID, assay, isolate, etc.) Thank you kindly.

@doremi No automation in the sense of robotic colony pickers or sample prep. It takes me roughly 30 minutes to spot 96 colonies onto a disposable MALDI target chip. The raw data is a peptide mass fingerprint (PMF), from which the software can generate IDs down to the species level, but only for organisms in the database. For colonies with no ID, there are dedicated Python packages for binning spectra into groups likely to be the same species. From there you could sequence as many isolates per bin as you deem necessary to confirm the identity of that PMF-type.

@MikeyC Got it. I'm hearing many folks warn about the limitation of the databases. That's  going to be a deal breaker for me since most of my target organisms are non-pathogenic and often only recently described. Thanks again.

Hard to know what you would consider "high-throughput", but it might be worth checking out the work by Kim Lewis at Northeastern University — see for example his 2020 Perspective in Cell1.
One of the tools his group uses is what they call the ichip, which allows them to grow "uncultivable" microbes2. This is done by placing the device in soil after sealing in 384 separate samples.
References:

Lewis, K. (2020). The science of antibiotic discovery. Cell, 181(1), 29-45.

Nichols, D., Cahoon, N., Trakhtenberg, E. M., Pham, L., Mehta, A., Belanger, A., ... & Epstein, S. (2010). Use of ichip for high-throughput in situ cultivation of “uncultivable” microbial species. Applied and environmental microbiology, 76(8), 2445-2450.


text:
What is this worm like animal found on a well water filter?
This was found in the water filter on a private water well that feeds a house in southeast Colorado, USA. The well has hard water, and is shallow (25’). There are several of these creatures all over the filter. They range from 3mm-5mm long.
The picture is of the water filter. In a video we have of them on the filter they move their arm like appendages.

Welcome to Biology.SE — this looks very interesting! I can think of a couple of marine invertebrates this vaguely resembles, but I have no useful idea about what it might be from well water. Can you [edit] in clearer pictures and/or possibly give a more detailed description of your creatures? Posting a link to your video might also be helpful. ——— Finally, it might be worthwhile to preserve specimens (e.g. stored in a clear high proof alcohol) — that way if no one comes up with a good answer you might be able to find a specialist at a college or university nearby who could identify them.


text:
What is the name of this strange white caterpillar?
My friend found this caterpillar in south-central Tennessee in August. No identification guides or databases I've searched had a similar picture. Does anyone know the name of this species?

Welcome to Biology.SE! For identification questions please [edit] your post to include the size of this creature. The size can be an essential clue for identifications. ——— If possible clear, closeup photos of the creature from multiple angles would also be helpful. ——— Please also take the [tour] and then go through the help pages starting with [Ask] questions effectively on this site. Thanks!

It is hard to be sure without clearer pictures, but that looks like something in the genus Eriocampa and it could be a Butternut Woollyworm (Eriocampa juglandis).
Picture taken in Kentucky from BugGuide.net © 2015 by dan40165:

These are the larvae of a sawfly (so not technically a caterpillar) and are about 1 inch long.
You can learn more about this particular species here. To narrow down the exact species it would be helpful to edit more information into your post including the types of plants in the area.


text:
Does gene editing have any technical cross-species limits?
How far could you go in cross-species gene editing?
Is it possible for example to introduce plant genes into human DNA and vice versa?
Question: How is gene editing limited by the "donor" and "target" species?

Putting plant cells into a human has nothing to with gene editing (and wouldn't work). I'd advise that you make clearer if you want to ask about this specific case (plants cells / genes into human) or about the general limits of gene editing.

thanks, of course cells come after all through gene experession, but fixed that now . Yes I want to know general limits of gen editing.

So, I'll assume that you are talking about gene editing in a laboratory (e.g. using a technique like CRISPR).
Theoretically, there are almost no limits to what you can do - or at least try. Fundamentally DNA is the same in all living (and non-living) things on earth, so you can absolutely take a gene from a plant and "paste" it into a human cell/genome (with some additional work needed to make the pasting work). You probably even go further and try to isolate whole chromosomes from one organism and (literally) inject them into cells of another one.
The problem with this radical approach to gene editing is, that it probably won't do much (good). A human cell likely doesn't recognise a plant gene as a gene and so just ignores it (it might still cause indirect problems for the cell though) and if you start making very drastic changes like adding chromosomes the chances that your cells will just die are super high.
This means that in practice you either need to (heavily) modify a gene you want to insert to match the structure and 'expectations' of the target organism - you basically have to make a humanised version of a given plant gene - or you can only use genes from similar enough organisms: as a general rule genes from the same kingdom (bacteria, archaea, plants, fungi, animals) are likely to retain some degree of function in another species (but how much exactly or even a general effect is almost impossible to predict).


text:
What does pseudo-irreversible mean?
In this publication (PDF) I encountered the following word:

Binding of rupatadine to histamine H1 receptors isolated from the
  guinea-pig cerebellum and lung was demonstrated by inhibition of
  3H-mepyramine binding; equilibrium inhibition constant (affinity) [Ki]
  values were 26–256 nM in the various experiments. Binding was
  time-dependent and pseudo-irreversible. Rupatadine was ~7.5 and
  10-times more potent than the structurally-related anti-histamines
  loratadine and fexofenadine. H1 receptor occupation by rupatadine in
  guinea-pig cerebellum and lung after oral (PO) dosing was relatively
  rapid (with maximum binding evident at 2–4 hours post-dose) and
  dose-dependent; low or no binding was seen at 48 hours post-dose.

In this context, what does pseudo-irreversible mean? Since it goes on to say the binding had dissipated within 48 hours, it seems to mean persistent or long-lived. Is that correct or does it have a more specific meaning in biology?

it seems to mean persistent or long-lived. Is that correct...?

Yes, it's correct, but how persistent the effect is is related to the rate of dissociation or rate of turnover of the receptor. It is relative, so to speak; relative to the effect of non-irreversible receptor binding.
Irreversible inhibition of, say, a cell surface receptor means that a drug (agonist/antagonist) binds permanently to the target receptor, usually by binding covalently; the chemical reaction is not reversible. This isn't synonymous with a permanent effect on a cell or organism; that depends on turnover of the receptor. A "faulty" receptor may be replaced quite quickly or not; it depends of the cell and the receptor.
Pseudo-irreversible means that the drug acts in a manner similar to  irreversibly bound agonists/antagonists, but there is no covalent bond formed; if a drug/molecule which has a sufficiently high affinity for the receptor is introduced (in a sufficiently high quantity), the new molecule will replace that which was strongly bound before (the pseudo-irreversibly bound drug.)
Some of these types of reactions have been studied more than others. One  example of this can be seen with h5-HT7 receptors, which are subject to pseudo-irreversible inactivation by risperidone and 9-OH-risperidone.
Clozapine and other competitive antagonists reactivate risperidone-inactivated h5-HT7 receptors: radioligand binding and functional evidence for GPCR homodimer protomer interactions

Sigh... why didn't you just answer the question in Health?

@CareyGregory - That wasn't asked on Health, at least not to my understanding.


text:
dNTP concentration for reverse transcription from RNA to cDNA
Does it affect the results if I use the half of dNTP amount in a reaction that it says in my protocol?
I usually use 4 microlitres per reaction and this time I did a mistake and I used 2.
The final cDNA will be used for qPCR relative cuantification,so I am not sure if that will affect my results.

It depends on several parameters like what is the concentration of dNTP you use, the length of the DNA amplified, the polymerase and other PCR parameters. I think the best way to know would be to check the quality of your result via agarose gel electrophoresis.

You will probably be fine. Lower dNTP levels might reduce the amount of product you obtained since it is usually the primers or the dNTPs that run out near the end of the reaction. The only other effect I am aware of is that the reaction might be slightly less specific as dNTPs sequester magnesium which is a big factor in determining the stringency of the reaction.
You say how many microliters you used but it would be more useful if you mentioned the concentrations of the dNTPs and the desired final concentration.

Please cite content that supports your answer, opinion based answers are bad.


text:
Tumour cell injection into a mice
I am going to inject mice tumour cells into mice to create them tumours and I am wondering if I need to have special precautions for that, even they are cells from mice tumour I am afraid what can happen if I inject myself by accident, I guess is not necessary to do it a hood either?

I hope you understand from the answers and comments that injecting mice (or any other thing) with live tumor cells is not without serious risks.

You should not be asking random people on the internet about this. Talk to your lab safety people. Get detailed instructions on handling requirements. As for the hood, again, do not ask here, ask your animal care people (IACUC or equivalent).  If you don't have both lab safety people and animal care people immediately available and involved before the experiment begins, you should not be doing this experiment.

And you will most likely need a permission by an ethics comission.

IACUC is an ethics commission. If humans are not being experimented on, there probably won't be a need for a second ethics review.

Ah, I didn't knew this abbreviation. But since I have seen people who wanted to start immediately, I thought I mention it briefly...

+1 This is occupational safety and health for labs 101: If you develop a procedure, it needs to be reviewed to ensure the proper personnel, training, controls and PPE are available. Before you perform the procedure!


text:
Has anyone confirmed Darwin's theory that nectar began as something "injurious" to sap?
In "Origin of Species", Darwin says (I have added bold for emphasis):

Certain plants excrete sweet juice, apparently for the sake of
  eliminating something injurious from the sap: this is effected, for
  instance, by glands at the base of the stipules in some Leguminosae,
  and at the backs of the leaves of the common laurel. This juice,
  though small in quantity, is greedily sought by insects; but their
  visits do not in any way benefit the plant. Now, let us suppose that
  the juice or nectar was excreted from the inside of the flowers of a
  certain number of plants of any species. Insects in seeking the nectar
  would get dusted with pollen, and would often transport it from one
  flower to another. The flowers of two distinct individuals of the same
  species would thus get crossed; and the act of crossing, as can be
  fully proved, gives rise to vigorous seedlings, which consequently
  would have the best chance of flourishing and surviving.

My first question is, has anyone done any experiments or chemical analysis or any other kind of science to show that nectar does in fact contain something "injurious" to the sap (or to the plant in general)?
More importantly, has anyone tried to "prove" Darwin's theory of the evolution of nectar in flowers?
I understand that Darwin seems to provide some hints by mentioning the

Leguminosae, and at the backs of the leaves of the common laurel.

as examples of a plant removing toxic substances from somewhere other than a flower, but I'm wondering if someone has "proved" that nectar in flowers started as a toxic substance the plant needed to remove.
Additionally, I'd be interested in how plants that haven't developed nectar/flowers deal with this waste (other than the common laurel and Leguminosae).  For example, does the plant just store the waste in a vacuole until it dies (please pardon my bad biology if "vacuole" isn't the correct organelle -- I'm not a biologist)?


text:
Are there fundamental differences between the adaptive immune systems of higher primates and other mammals?
I recently attended a Leukemia and Lymphoma Society lecture in which one of the lecturers indicated that there are fundamental differences between the adaptive immune systems of higher primates and other animals.
The Wikipedia article on adaptive immune system seems to indicate that mammals share very similar adaptive immune systems.
I understand that there are (minor) differences between the B cells and T cells (and other cells) of different species, and that two species can differ in their susceptibility to a particular kind of pathogen (virus, bacteria, whatever), depending on small differences in cell surface proteins, etc.  I am not interested in these kinds of small differences.
Rather, I would like to know if there are any fundamental differences between the adaptive immune systems of humans (and other higher primates) as compared to other mammals.  For instance, are there particular kinds of adaptive immune cells in primates that don't appear in other mammals.  Or are there fundamentally different mechanisms (e.g. the mechanism of T-cells activating B-cells) or lymphatic system organs that are markedly different in higher primates as compared to other mammals.
In short, are there big differences in the immune system of higher primates that make them stand apart from the other mammals in the same dramatic way that thumbs and intelligence seem to separate them from other species?

When the lecturer said other animals, were they specifically referring to other mammals or did they just say other animals? If other animals that can mean everything from fish, jelly fish, sponges, corals, reptiles, birds, insects, worms, etc., so then yes there are differences. If they specifically said other mammals, then there may be subtle differences, but in research animals such as mice, it is very similar. They may have meant the constant regions of antibodies, as these are unique to the species and can actually be exploited in immune assays.

In fact, it is often referred to as the vertebrate immune system. This might help: http://www.ncbi.nlm.nih.gov/books/NBK2388/

I am not aware of a discriminatory feature between primates and all other mammals. However, there are significant differences for some animals. For example, the anti bodies of camelids and sharks consist of the heavy chain only, or the antibodies of sea lampreys are not of the immunoglobulin protein family.
Those differences have been exploited for biotechnology.
consider this review: http://www.nature.com/nbt/journal/v23/n10/full/nbt1127.html


text:
Do archaea undergo the same horizontal gene transfer processes as bacteria?
Do archaea also undergo processes like conjugation, trasformation, etc.? If not, do they have their own horizontal gene transfer methods? Can bacteria conjugate with archaea?


text:
How to interpret Glimmer scores?
I am trying to find orfs with Glimmer. At the end, Glimmer gives me a .predict file with the orfs and a corresponding raw score.
When I look at the documentation, it says about the score, "This is 100 times the per-base log-odds ratio of the in-frame coding ICM score to the independent (i.e., non-coding) model score. It gives a rough quantification to how well an orf scores that can be compared between any two orfs." You can look here for the documentation-
https://ccb.jhu.edu/software/glimmer/glim302notes.pdf
But what does the score really mean in an absolute manner? How do I realize for an orf, if the score is significant or not?

The score is the log odds ratio, times 100, that the ORF that you input is a coding region. For example, if Glimmer thinks the log odds are 10 to 1 that your region is coding, then your score will be 10x100 = 10000. Log odds of 10:1 imply more than a million to one certainty.

Thanks. But the documentation says log-odds ratio. But even then, my question is what is a good odds ratio absolutely. More abstractly, when can I be confident that this orf is really a coding region.

You can't really be sure without experimentation and research, because glimmer can screw up. But a score of 300 or more suggests better than 90% odds that it is a coding region according to Glimmer. 10000+ implies a million to one odds.


text:
Proteins that cannot form biofilm?
I am trying to build a machine learning training set for bacterial protein sequences that form biofilm, and that cannot. I collected the positive sequences from the GO ontology website but for negative sequences I am not sure which sequences to incorporate into my training set.
Can anyone point me to resources for proteins sequences that are known to be not capable of forming biofilms?


text:
DNA to Binary Distance Computation
If I represent DNA as binary values, what is the best way of computing distance between them.
So :
A = 00,
T = 11,
G = 01 and
C = 10
Hamming Distance between ATGC and TAAC is 3, however their binary representations give a different answer:
Hamming Distance of 00110110 and 11000010 = 5.
Whats the best way of distance computation if the DNA bases are represented in this way?

It is a question of theoretical computer science, not biology. I am voting to close. You should give it a try on cstheory.SE.

Agree with @Remi.b. But before you leave us, "Why would you want to do that?" as IT Support used to say.

Ask this question on StackOverflow not cstheory

I found a solution, I'll answer it when you ask it again on StackOverflow

This is a relevant question for biology.se, I think. "what is the best way of computing ..." phrase in the question is misleading, however. The question is not about how to perform a computation, but rather how to represent some biological entity formally, in a biologically meaningful way. This is a question about theoretical biology, not about cs.

I encountered this problem when studying M.A.Nowak's book called Evolutionary Dynamics. There, the author introduces the concept of a sequence space, where he attributes "imagination" of it to John Maynard Smith.
Nowak's discussion is very superficial and does not include your question, and I don't have an answer to it, either.
But i think I can make the question a bit clearer.

A sequence space is a L-dimensional space for sequences (of DNA or whatever) of length L. Each sequence is a point in that space, and position of the point is determined such that, value of each position within sequence determines the coordinate of the corresponding dimension in the sequence space. Author suggests then the Hamming distance of sequences within this sequence space as a metric for similarity between sequences.

These definitions make perfect sense, when the sequences have binary elements. And also any sequence can be represented by a corresponding binary sequence, says the author, implying that the rest is straight forward, if we want to extend the definitions to any biological sequence.
But this is not a straight forward task, as the example in the question demonstrates.

Each coordinate, or position, in the sequence will be represented by 2 bits. But some of the bases differ in one of their bits (A and G for example), whereas some of them (A and T) differ in both of their bits. Of course plenty of speculations can be done about how to do it conveniently, but there is no obvious way, as far as I can see. A good answer could provide some examples from the evolutionary dynamics literature for example, about how it is done.

A last note, this more of a evolutionary dynamics / theoretical biology question, than a bioinformatics question. It is relevant, when a formal representation of biological sequences (mostly for simulation purposes) is needed. For more practical purposes, about calculating distance between actual sequences, Jack Aidley's answer provides some good foundation.

Thanks for telling me why you want to know. I imagine that comparing two alphabetic strings is not the same as comparing two binary strings even though you have 'converted' one to another. Which should one use for evolutionary comparison? No idea! To answer that I'd need to understand the Hanning distance to decide whether or not it is applicable to the problem. So I still think you need a computing scientist with some molecular genetics to discuss this with.

I didn't ask the question by the way.

The best way is to choose a distance that represents what you want it to rather than necessarily relying on the Hamming distance.
If you simply want base-by-base difference then calculate that (this may help) but you may also want a difference that depends on the likelyhood of mutating between different bases in which case you want to define a function that translates the mutation into a score for each transfer, i.e. you might want to score a deamination from 5-methylcytosine to thymine as the most likely occurrence. Expressing the relative likelyhoods of different mutations is not an easy problem but there are a number of widely used options.
The important thing is to ensure you represent the underlying biology not ensure that you have the fastest implementation. Decide on this first, then on the algorithm that gives you the best speed (also, deciding on that algorithm is on-topic for Stack Overflow not this Stack Exchange).

I am reluctant to answer with code, but it seems that the community decided that was an appropriate question for Biology.SE. So here is my solution.
The idea is to "compress" the two bit that represent each nucleotide, such that each nucleotide will contribute 0 or 1 (not more) to the distance.
You could use binary operations to do something like this (in Java, but you can apply the logic in any language):
int seq1 = 54, seq2 = 194;//ATGC and TAAC
int evenBit = 0xAAAAAAAA, oddBit = 0x55555555;

int pseudoDist = seq1 ^ seq2; //Integer.bitCount(pseudoDist) is 5
int dist = ( (pseudoDist&evenBit)>>1 ) | (pseudoDist&oddBit);
int finalDist = Integer.bitCount(dist);//output 3 not five

The idea is to get the total number of bits that are different with:
seq1 ^ seq2

But you can't just count the bits yet, because you will get the hamming distance instead, so you have to compress all the bits that correspond to the same nucleotide to the same bit using: (pseudoDist&0xAAAAAAAA>>1) and pseudoDist&0x55555555. The first one keeps the bits on even positions and the second the ones on odd positions.
Now you use evenBits | oddBits, and you can count the bits.

The original question conflates mathematical operations with the edit distance between two strings. A hamming distance is a measure of the number of changes you have to make to convert one string into another string. Converting the the alphabet into binary digits and then adding or subtracting the numbers is not going to tell you how many edits were required.

@mdperry The fact that you don't understand it, does not invalidate it... You say it's impossible, but have you looked at my answer, have you tested it? Look it says that the hamming distance between ATGC and TAAC is 3, which is the right answer.

If your approach and solution is correct then it should be straightforward to apply your code to the examples on this page: https://en.m.wikipedia.org/wiki/Hamming_distance.

@mdperry this is not meant for any string, the fact that there are only four states possible allows you simplify it into 3 lines of code, that works for this particular problem

Actually, I now agree with you: my comment is incorrect, my apologies.

This encoding doesn't make sense as nucleotides are not in the Hamming space. Hamming distance between every two nucleotides is constantly 1, but in binary encoding, it varies from 1 to 2.


text:
How to improve A260/230 ratio?
I am extracting DNA from cicada exuviae using ethanol precipitation combined with either NaCl 6M or Ammonium acetate. After DNA extraction, my samples have quite low A260/230 ratio such as in the attached picture. I suspect this might be due to left-over salt in my samples, since I dont use kits or phenol/chloroform/isoamyl in DNA extraction protocol. I proceeded on with PCR using 16S primers but received no bands.
Could you advice what potential contaminants could appear in the samples that prohibit PCR? As well as what could I do to improve this ratio? Thank you so much!

Are the cultured cells you use in the beginning or tissues? How do you break your cells?

Actually, I used exokeleton, which contains left-over tissues of cicada when it molts.
For the lysis, I used 600 ul buffer including Tris 10 mM, pH 7.5, NaCl 400 mM, EDTA 100 mM, and SDS 0.6%, along with 30 ul Proteinase-K.


text:
Do symptoms of common cold infections correlate with the infectious agent?
The common cold - in summary, an acute upper respiratory tract viral infection - is one of the most frequent viral infections in humans and can be caused by about 200 viral types [Eccles (2005)]. Symptoms are thought to be caused by the immune response in the host, rather than by actual damage caused by the virus [(Hendley (1998)].
My question: is there a correlation between the symptoms observed during a common cold and the viral type that causes the infection? Of course, one can find out the cause of the infection diagnostically by taking samples and isolating DNA or setting up cultures, but that is not my question. My question is whether it is possible to predict the virus taxon causing the infection by symptoms.
My prediction from the information I have at hand is that it should, at least in tendency, be possible as different infections agents use different infection pathways that in return invoke different immune responses and, finally, symptoms [Eccles and Weber, ed. p. 107 - 147 (2009)]. However, I found a rather dated article by Tyrrell et al. (1993) that analyses infections caused by several rhinovirus strains, a coronavirus, and of the respiratory syncytial virus (virus taxa among the most common infectious agents for common colds). They conclude that the main difference between colds induced by different viruses is in duration of the incubation period, whereas patterns of symptom development were not substantially different with different viruses. So is this the current state-of-the-art?

The strains can mean the difference between mild or severe symptoms though (just think spanish flu vs H1N1). Symptoms can also be different depending on the person infected (some have headaches, some don't) since it depends alot on how your body reacts to the virus, I would think that differentiating viruses by the symptoms would be extremely hard, but severe symptoms could suggest some specific strains for example.

Not going to post as an answer because finding the literature to support it would be too time consuming. No, symptoms are inadequate to differentiate among viral agents with a few rare exceptions. There is a dearth of research because it doesn't really matter, does it? There is no treatment for the vast majority of respiratory viruses, and more importantly, the illness is for the most part mild and self-limiting. To fund research, there has to be a fairly good reason besides curiosity (like ability to treat/prevent, or infection causes significant morbidity/mortality, e.g. RSV/H1N1.)

You cannot necessarily identify the cause of a particular case of the common cold by culture or nucleic acid tests. These tests will tell you if an organism is present, not if it is causing disease, and many of these organisms are present in samples from individuals who do not have a cold.

@anongoodnurse I think the dearth of research is more the result of lack of interest than lack of a reason. The common cold is one of the most common causes of provider visits and causes substantial economic burden on a population level. The treatment is simple and appears to be effective: empathy, reassurance, and counseling about appropriate otc remedies for symptom relief.

@DeNovo - "the dearth of research is more the result of lack of interest..." There is no lack of interest (or research) in the common cold; as you said, it is a very significant societal burden. What there is no interest in is correlating the very, very common symptoms with which of the hundreds of possible viruses may be the cause.

@anongoodnurse this is true, but i'm sure you wouldn't be surprised to hear that residents fall asleep as soon as I start talking about how to approach the common cold.

@DeNovo - lol, residents fall asleep the moment they stop moving. As a resident, I fell asleep listening to a patient's heart. I've heard more embarrassing falling-asleep stories. :) That said, the common cold is bread and butter; they should be listening.


text:
How many (taxonomic) families are there?
Approximately, how many families have been identified?
I've often often come across figures for the total number of species on Earth. Recently, I found myself wondering about the encompassing ranks above them, specifically family, but I can't recall any figures on family and it's a little difficult finding what I want by typing "family" into a Google search. ;)

Families (and really, any taxa except species, to some extent) are artificial, fluid and ever-changing. That's not to say they aren't useful, but the number of families is likely to be changing quite a lot on a nearly daily basis, as more and more families are converted into monophyletic groups based on DNA evidence.

... but just as a round estimate its thousands at least.  browsing encyclopedia of life I see scores just in the flowering plants...

This was asked three years ago. I am curious what the numbers look like today.

Not so many they couldn't fit on a very big boat.

The 2011 paper How Many Species Are There on Earth and in the Ocean? indirectly answers this question as well as any other source you'll find I imagine. It estimates how many species there are total based on the rate of discovery of higher taxa; it includes plots of number of taxa over time for the major groups of life in Figure S1. Which gives:
Animalia - 5300 families in 2011 (the plots are given with only one significant figure, the second one's my estimate) (estimated total: 5800)
Chromista - 270 families (estimated total: 360)
Fungi - 550 families (estimated total: 620)
Plantae - 750 families (estimated total: 800)
Protozoa - 280 families (estimated total: 310)
Archaea - 27 families (no estimated total; the number has been increasing exponentially so far)
Bacteria - 300 families (same as for Archaea)
Which gives us a total of 7477 families in 2011, with an estimated total of (ignoring Archaea and Bacteria, who don't really fall in the same kind of classification anyway) 7890 families. (make that 7500 families discovered by 2011 and 8000 estimated in total given the imprecision involved in my reading the plots).

The NCBI Taxonomy statistics page displays the following information:
There are currently 73540 genera, 331418 species, and 23127 taxa of higher order. Since the number of taxa decreases with the genericity of the taxon, there are probably around 20000 families, give or take a few thousand.

The Plant List has 642 families listed: http://www.theplantlist.org/1.1/about/#changes.
For a quick comparison, Wikipedia lists

522 fish families:
en.wikipedia.org/wiki/List_of_fish_families
136 mammal families:
en.wikipedia.org/wiki/Mammal_classification
61 amphibian
families: en.wikipedia.org/wiki/List_of_amphibians
57 reptile families: en.wikipedia.org/wiki/List_of_reptiles.

This only covers a subset of the total, however (see here for other animal classes). It is relatively difficult to find a good, and relatively complete answer to this, though.

I don't know about other groups, but about plants, number of families depends on the system you follow. Recent version of The Plant List (1.1) estimates about 352 000 species of Angiosperms and lists over 400 families. See http://www.theplantlist.org/1.1/browse/A/ It is very good and reliable source of information. Second very good source about plant systematics is database Tropicos provided by Missouri Botanical Garden and Angiosperm Phylogeny Group. See list of families according to APG3 http://www.mobot.org/MOBOT/research/APweb/ and click to "Families" on the top of the page. Just note, that there are also synonyms, so that You can't just count the rows. ;-)

I'm adding this as another answer since it's a different source, but Gaurav gave this excellent resource as an answer to this related question:

The Integrated Taxonomic Information System (ITIS) is maintained regularly by a consortium of North American governmental agencies, and will give you a list of all the classes, orders, families or genera in any of seven kingdoms they recognize. It might be biased towards North American taxa, but it might be quicker to get a list from then rather than extracting them from Wikipedia.


text:
Are the subordinate genes of a repressed operon really "turned off"?
Operons are often described using all or nothing language. A repressor binding to the operator is usually presented as "turning off" the regulated genes. Case in point, Scitable at Nature.com says:

In addition to being physically close in the genome, these genes are regulated such that they are all turned on or off together. ... The ability to turn ... genes on or off as a group therefore provides an efficient way to quickly adapt to environmental changes. ... mutations affecting the promoter can prevent all of the operon's genes from being expressed, ...

The thing that confuses me is that I've also heard genetics aren't all or nothing, that genes can only be "down regulated" or "up regulated", not truly turned on and off.
Obviously, differing amounts of moderate levels of whatever effector is modulating the repressor could lead to different levels of downstream expression. But, what about situations where nothing is inactivating the repressors? Will the regulated genes be (for all intents and purposes) be turned off? Additionally (and relatedly), is it even a realistic scenario for an operator to always (or almost always) be bound to by such a repressor?

There are several mechanisms by which the expression of a gene can be completely turned off. Certain network architectures can ensure foolproof repression (for e.g. by using multiple repressors parallely or additional epigenetic silencing mechanisms). Bistable switches can also ensure robustness of expression in a way that small fluctuations in the transcription factor levels do not lead to expression of the gene. Sometimes the activator molecules need to co-operatively activate the expression. In such conditions the activation graph is sigmoidal with respect to activator concentration. All these mechanisms can prevent leaky expression.
Some genes do have a basal expression but it is not necessary for all cases. However, basal expression is absolutely necessary for proper functioning of the lac operon (see this post).
Having said this, there is always some non-zero probability of expression of a gene when it is actually turned off. This probability may be very low but it would not be zero. This small accidental expression may or may not have any significant downstream effects.


text:
What makes DNA sequences most different/recognizable from a biological perspective?
We can pretty easily quantify the amount difference between two different strings/sequences of characters. For example, if we take the words trebuchet and trebucket, we can say they have a Levenshtein distance of 1 (only one character-edit's worth of difference).
From a linguistic perspective, that distance is very noticeable, whereas the difference between compliment and complement is much less so. Even though both pairs are only one edit apart (respectively), not all differences are equally distinguishable.
What is the biological equivalent of this when it comes to DNA binding proteins and RNAs? How do I identify which DNA sequences are more recognizably distinct from each other? For example, if we took a DNA binding protein that recognizes the sequence TGCCTCGAA, is it more likely to misrecognize AGCCTGGAA than TGCCAGGAA (or vice versa) as its target sequence?

That's going to vary from one nucleic acid binding protein to another. Remember, DNA and RNA are not simply  sequences of letters, but there can be secondary structure, as well as more basic chemical interactions between adjacent or more distant bases - see RNA hairpins as an example. All of this gives a "shape" to the recognized portion of nucleic acid, which can inform binding capacity just as much as, or potentially even more than primary sequence alone.

@MattDMo That is certainly not surprising, but it would seem to me that there should be some parameter to that variation. For example, perhaps switching a single base in a sequence from a purine to pyrimidines is more distinguishable than to the other purine.

Related: http://biology.stackexchange.com/q/38808/3340

This isn't a question with a really well accepted answer yet, and comes up quite a lot in e.g. studies of population variation in transcription factor motifs.
Usually, we approximate the sequence preferences of a DNA-binding protein with a position weight matrix. A weight matrix will given you a score for two sequences,  so the simplest means of quantifying the relative binding strengths for two sequences is compare these scores. You could also, say, compare the relative chances of getting the scores under some background distribution of scores, which would arguably be more comparable between different factors.
The PWM score (and other means of describing sequence specificity) are only an approximation to what you're really interested though, which is the binding energy associated with the interaction. If you have detailed experimental evidence about how the protein binds, you might use that, and there are some papers that build models for approximating it from PWM scores as well. However the reality is that you can only ever roughly approximate what's happening in the cell, because of molecular crowding, and the many other factors that will be present on the chromatin fibre, so the difference in PWM scores is often what gets used.
Predicting the impact of a given change to a non coding DNA sequence is a hard problem. Sometimes you'll get a decent approximation, but in the end you're trying to do biochemistry from first principles, and we aren't there yet.

DNA is a chemical, and therefore, its interactions are governed by its shape.  There is no way to look at a DNA sequence and know all the ramifications that changing a letter will have on its shape.  I could tell you that changing the first two or last two letters of an intron are highly likely to destroy a splice site, but you can't make hard and fast predictions about DNA binding in many other situations.


text:
How much time do the different mechanisms of gene regulation need to take effect?
I am thinking of the major regulatory mechanisms, like general transcription factors, activators, repressors, and RNA interference.
If non-active regulator genes using each of the different mechanisms were activated, what would be delay (on average) between each of those genes' activation and their downstream regulatory effect?

I don't have an answer for you, but people often imprecisely say that type of regulation takes place on the order of many minutes to hours, and that may be as precise as you can get given the variable kinetics of any given pathway. I will say this though: the kinetics of knockdown in RNAi could largely depend on the rate of degradation of the protein (which is HIGHLY variable). Even if you take away the mRNA, the protein is still there until it's degraded. If it's normally degraded fast, the kinetics of mRNA degradation will be limiting. If it's slow, then you have to wait for the protein.

Also, all genes in eukaryotes require general transcription factors, but basically all of them also require activators to act first to recruit the GTFs and histone modifying components (which allow GTFs and their friend RNA polymerase to access genes). So, they are really downstream of activators and  their recruitment is part of what contributes to the kinetics.

People often imprecisely say that type of regulation takes place on the order of many minutes to hours, and that may be as precise as you can get given the variable kinetics of any given pathway.
Also, all genes in eukaryotes require general transcription factors, but basically all of them also require activators to act first to recruit the GTFs and histone modifying components (which allow GTFs and their friend RNA polymerase to access genes). So, they are generally downstream of activators and their recruitment is part of what contributes to the kinetics in euakryotes.
For RNAi, the kinetics of knockdown in RNAi could largely depend on the rate of degradation of the protein (which is HIGHLY variable). Even if you take away the mRNA, the protein is still there until it's degraded. If it's normally degraded fast, the kinetics of mRNA degradation will be limiting. If it's slow, then you have to wait for the protein to be degraded by its natural mechanisms (https://en.wikipedia.org/wiki/Proteasome).
Sigma is saying RNAi effects can be seen in around 24 hours
http://www.sigmaaldrich.com/life-science/functional-genomics-and-rnai/sirna/learning-center/mission-sup-reg0/experimental-design0.html#s3
Thermo is implying something similar by saying RNAi effects can be seen for 5-7 days post-transfection (i.e. they imply that it can be seen after a day).
https://www.thermofisher.com/ca/en/home/references/ambion-tech-support/rnai-sirna/tech-notes/duration-of-sirna-induced-silencing.html


text:
Are bacterial RNA polymerases distributed evenly throughout the cell or localized?
Bacterial cells aren't internally compartmentalized with membranes (like eukaryotes). This naturally leads to an image of a homogeneous interior, but bacterial cytoplasm isn't homogeneous. Case in point, the DNA is localized in the nucleoid. If the cell goes through the trouble of putting all its DNA in one spot, having RNA polymerases everywhere would be pretty wasteful (even counterproductive). So, is the more intuitive image of bacteria, with RNAPols everywhere, right? Or are they localized? If so, how?
If species matters, lets primarily talk E. coli.

In E. coli most RNA polymerase molecules are bound to DNA, and those that aren't are within easy reach of the DNA.
According to Larry Moran

A typical E. coli cell contains about 5000 molecules of RNA polymerase. When the cells are growing rapidly, 2500 molecules will be bound to genes in transcription complexes. Another 1250 will be in initiation complexes of various sorts and most of the remaining RNA polymerase molecules (1200) will be bound to DNA non-specifically. Only a small number (~50) will be free in the cytoplasm.

The numbers that he quotes for number of molecules of polymerase and proportion that are engaged in transcription are in reasonable agreement with values available at Bionumbers. So it seems that most RNA polymerase molecules will be located in the nucleoid; any molecules that are free in the cytoplasm are within easy reach of finding a DNA binding site since a typical protein can diffuse across an entire E. coli cell in 10 ms. (Bionumbers)


text:
Can we measure the rate of mutation in (say) E. coli without gel electrophoresis or genome sequencing?
If so, how? And, what are the pros and cons?

This is pretty broad question so I'm not really sure what the end goal is here, but here goes.
The answer is both: "yes" and "it depends what you mean exactly". I'll explain what I mean.
Firstly, you should clarify in your question whether what you actually mean is the substitution rate, or not. The mutation rate, which is how often a given base pair position changes, cannot be observed with any current technique, even Next Generation Sequencing. This is for 2 reasons (that spring to mind, there may be more).

If you make an observation at time $t$, and another at time $t +1$, but a base mutates (say A $\rightarrow$ C) and then mutates back again between your observations, you would not be able to tell that this had ever happened. Thus there would be 2 mutation events which you would not count in your 'rate'. This is one of the reasons molecular clocks based on SNPs tend to underestimate ancestral distances.
If a mutation occurs that is deleterious (has a negative effect on the organism), the mutation won't be observed because the lineage of the organism won't survive to be sampled. By the way, the chances of a mutation being deleterious are considerably higher than it being advantageous, but lower than it being neutral.

The substitution rate is how many mutations have occurred that persist in the lineage. Now if we assume that when you say mutation rate, you mean substitution rate...

Yes, you can study mutation rate without 'molecular techniques'.
Throughout the history of classical genetics we've studied mutation rate without access to these tools.
My favourite example is basically the one I used in this question, where you can study the emergence of antibiotic resistance in bacteria (since you provided the example of E. coli, this is rather apt).
In the papers I linked however, they will almost certainly have used some molecular techniques to validate their answers.
But, if we set that aside for the moment, the very first indication of antibiotic resistance one might observe in the lab is seeing colonies of a bacteria growing on a petri dish in the presence of an antibiotic they are supposedly susceptible to.
Now, if we consider the 'mutation rate' to be the time taken (perhaps counted in the number of generations), for a spontaneous mutation to appear as a phenotype (observed characteristic), you can assess this purely visually without any molecular techniques. If you repeated this experiment however, you would most likely not be able to verify that resistance arose in exactly the same way twice without use of various molecular technqiues.
As a molecular biologist though, I would not really consider this a very good approach/definition of mutation rate, which leads me on to...

'It depends what you mean exactly'
If you were to stick to what is probably the stricter but more widely accepted definition of mutation rate (.i.e substitution rate), you'd need molecular techniques to determine this, as you'd need to know precisely which bases are changing.
Hand-wavy list of pros and cons:
Phenotyping (pros and cons):

Quick
Cheap
Easy
Low resolution
Possibly not reproducible

Genotyping (pros and cons):

Massive resolution
Can be quick in some circumstances
Functional information
Generally expensive
Requires specialist equipment

Not an exhaustive list by any means.

Here is a classic reference that shows how you can use bacterial mutation rates to test compounds to see if they are carcinogens:
https://www.ncbi.nlm.nih.gov/pubmed/4151811


text:
Why are high levels of epinephrine and low levels of cortisol signs of stress?
According to my psychology textbook (Psychological Science: Modeling Scientific Literacy), high levels of epinephrine and low levels of cortisol are signs of stress. However, earlier in the textbook it is shown that the autonomic response of the human body releases norepinephrine and epinephrine, while the hypothalamic pituitary adrenal (HPA) axis pathway releases cortisol. So it seems to me that signs of stress would be high levels of both of these things. Is my textbook wrong or am I just understanding what stage of stress it's describing?

So it seems to me that signs of stress would be high levels of both of these things.

Yes. Wikipedia claims [1] the same:

Stressors [...] activate the HPA axis, though via different pathways. [...] Stressors that are uncontrollable, threaten physical integrity, or involve trauma tend to have a high, flat diurnal profile of cortisol release (with lower-than-normal levels of cortisol in the morning and higher-than-normal levels in the evening) resulting in a high overall level of daily cortisol release. On the other hand, controllable stressors tend to produce higher-than-normal morning cortisol.

The same on a webpage of Society for Endocrinology [2]:

In addition, in response to stress, extra cortisol is released to help the body to respond appropriately.

Yet, there are conditions in which stress can lower cortisol levels. One of them is psoriazis [3]:

[...] patients with persistently high levels of stressors seem to have a specific psychophysiological profile of lowered cortisol levels and may be particularly vulnerable to the influence of stressors on their psoriasis.

And some psychiatric disorders [4]:

Several stress-associated neuropsychiatric disorders, notably posttraumatic stress disorder and chronic pain and fatigue syndromes, paradoxically exhibit somewhat low plasma levels of the stress hormone cortisol.

References:

Wikipedia contributors, "Hypothalamic–pituitary–adrenal axis," Wikipedia, The Free Encyclopedia, http://en.wikipedia.org/w/index.php?title=Hypothalamic%E2%80%93pituitary%E2%80%93adrenal_axis&oldid=619115237 (accessed August 7, 2014).
Society for Endocrinology. Hormones. Cortisol (2013). Available from http://www.yourhormones.info/hormones/cortisol.aspx (accessed 07.08.2014)
Evers AW, Verhoeven EW, Kraaimaat FW, de Jong EM, de Brouwer SJ, Schalkwijk J, Sweep FC, van de Kerkhof PC. How stress gets under the skin: cortisol and stress reactivity in psoriasis. Br. J. Dermatol. 2010 Nov;163(5):986-91. doi: 10.1111/j.1365-2133.2010.09984.x. PubMed PMID: 20716227.
Yehuda R, Seckl J. Minireview: Stress-related psychiatric disorders with low cortisol levels: a metabolic hypothesis. Endocrinology. 2011 Dec;152(12):4496-503. doi: 10.1210/en.2011-1218. PubMed PMID: 21971152.


text:
Is there a difference in the energy produced from the hydrolysis of GTP and ATP?
Is the energy produced from the hydrolysis of GTP greater than/less than or the same. I know that GTP can be converted to ATP through a pathway and so my thinking was that GTP would be less.

The base attached to ribose has, I guess, negligible effect on the stability of the third phosphate. So, I guess the energy released should be the same. Still looking for valid references.

As @WYSIWYG said in comments, there is no effect of change in nitrogenous base on the third phosphate. Why? Simple, they are too far to directly influence each other's stability. See the diagram (of ATP) below from wikipedia:

and of GTP from here:

What one can clearly interpret from these diagrams is that there would be no effect on the energy released by hydrolysis of the third phosphate by the nitogenous base present. It will remain -30.5 kJ/mol(1) in both cases.

You are almost certainly correct, but you do not cite an experimental value of the free energy of hydrolysis of GTP to support your statement, which I think is required. (And not so easy to find.)

You are correct, there is (probably) no citation for it. I'll  search for it tomorrow when I'll  be free :)


text:
why can proteolysis only occur once on a enzyme?
My texbook says that proteolysis can occur once in the lifetime of the enzyme? I know that proteases must cleave the zymogen at more or more specific locations, but why can't the process take place more than once?

Because you mention zymogens, I assume the proteolysis you are referring to is the conversion of the inactive to active form of an enzyme by cleavage. The reason this would only happen once is that there isn't any process to put the two cleaved halves of the protein back together, so once cut in half it stays that way until degradation.
The proteolytic enzyme that is doing the cleavage of the zymogen will repeat this process over and over with multiple copies of the zymogen: it is the zymogen itself, the target, for which this is a once-in-a-lifetime event.
It would certainly be possible for a zymogen to require multiple cleavage events, but each of those events could only happen once.

Just to clarify - proteolysis at a particular site in a zymogen only occurs one in the lifetime of the zymogen. The protease can cleave many many many individual zymogen proteins within its life.

@MattDMo Yes indeed - maybe this is the confusion the question asker had in the first place, I will add to my answer.


text:
What is the difference between endometrial lining and endometrial stroma?
This is in the context of embryology whereby the syncytiotrophoblast erodes the endometrial stroma during the penetration and implantation of the zygote.
But I don't know the correct terminology of what exactly has been eroded.