Produced by Loredana Manfra of Institute for environmental protection and research (ISPRA-Department of Environmental Quality Monitoring) and Federica Savorelli of Regional Agency for Environmental Protection (Emilia-Romagna, Italy).
Reference Members for the sub-working Group on the Artemia crustacean within the Biological Methods Group &#x2013; Saltwater/Brackish Water and Sediment of the UNICHIM Commission on Water Quality (the institute for technical standardization operating in the chemical sector).
Videography and editing by Marco Pisapia of ISPRA (Web Unit). 
Production Coordination by Anna Maria Cicero and Erika Magaletti of ISPRA (Department of Environmental Quality Monitoring).
We would like to thank the Regional Agency for Environmental Protection of Emilia-Romagna, Ferrara Branch, the Biological Methods Group - Saltwater /Brackish Water and Sediment of the UNICHIM Commission on Water Quality and Luciana Migliore, Tor Vergata University (Rome) for their cooperation; Rossella Boscolo and Massimo Gabellini of ISPRA (Department of Prevention and Mitigation of Impacts) for their financial support; Fabio Matassa and the London School of Languages in Rome for their linguistic support.

1. Method Standard 
Our testing involves exposing Artemia larvae to an interval of substance concentrations or a test sample for the purpose of determining the concentration or dilution levels that would cause the death of 50% of the organisms (LC50) exposed over 14 days and meeting the conditions defined by this method. If necessary and possible, the following may also be determined: a) the concentration level that causes the death of 20 % of organisms exposed (LC20); b) the highest analyzed concentration level that does not determine a higher mortality rate than that of the negative control (NOEC); the lowest tested concentration level that determines, after 14 days, a higher mortality rate than that of the negative control (LOEC).
2. Testing Materials 
2.1 Artemia eggs 
To perform testing, Artemia franciscana larvae of less than 48 hours old (nauplii) are required, at stage II or III, obtained from newly hatched eggs5. Certified eggs or cysts utilized to support testing are commercially available, e.g. from the Quality Assurance Research Division, U.S. Environmental Protection Agency, Cincinnati OH 45268, U.S.A. or from the Laboratory for Biological Research in Aquatic Pollution, University of Ghent, Belgium. 
2.2 Water dilution
Synthetic saltwater may be used as dilution water, prepared by dissolving recognized analytical quality reagents or a commercially available formulation in distilled or de-ionized water. Salt blends for creating the ideal saltwater are commercially available, e.g. the Instant Ocean mixtures. It is recommended a dilution water with a salinity equal to 35 (&plusmn;2) PSU as well as a dissolved oxygen concentration of more than 80% of the value of saturation and a temperature of 25 (&plusmn;2) &deg;C. After aeration for 48 hours and filtration in filters with a porosity of at least 0.45 &mu;m, the dilution may be stored for a maximum of 30 days in the dark at a temperature of 0-4 &deg;C. 
2.3 Algal culture
It is recommended that test organisms are feed on Dunaliella tertiolecta microalgae, which grow exponentially and reach densities of 1.3x106 - 2.0x106 cells/ml. Testing on environmental samples requires cultures with densities higher than 2.0x106 cells/ml6. It is suggested that Dunaliella tertiolecta grows in a culture media containing seawater (30 (&plusmn;2) PSU and 20 (&plusmn;1) &deg;C) with addition of essential nutrients, minerals and vitamins. Some indications on the algal media composition and preparation are reported in the Appendix. The initial cell density is suggested to be approximately 105 cells/ml. The algal culture is maintained in a room thermostat at 20 (&plusmn; 1) &deg;C, illuminated with fluorescent lamps 3000 (&plusmn; 300) lx, with a photoperiod ratio of 16 hours of light to 8 hours of darkness.
Other phytoplankton species could also be used for feeding but it would be necessary to make comparative studies with these phytoplankton species to find the "equivalent" species to Dunaliella tertiolecta, because the experimental conditions (e.g. initial cell densities, light illumination, temperature, composition of culture media and exposure time) varies among phytoplankton species. Besides the tolerance of the microalgae to different pollutants needs to be considered because the pollutant effects on the growth and development of phytoplankton impact also to the Artemia.
2.4 Reference substance
The reference substance in this test is Sodium Dodecyl Sulfate.
2.5 Laboratory glassware
In addition to the usual laboratory equipment, the following material is needed:
<ol>
 <li> Testing containers including 100 ml borosilicate glass beakers (18 to perform testing and 18 for transferring solutions).
 </li>
 <li> A flask (e.g. 500 ml flask) to prepare the standard solution. </li>
 <li> Six flasks (e.g. 200 ml flasks) to prepare the testing solutions. </li>
 <li> Glass or polystyrene 5 cm diameter Petri dishes with removable covers to be used for activating the cysts and transferring Artemia from the hatching dish to the testing containers. </li>
 <li> Glass Pasteur pipettes (with tips rounded by flame) for transferring the nauplii. </li>
 <li> Glass cannulae or Pasteur 3 ml disposable plastic pipettes (to be cut) for transferring Artemia. </li>
 <li> Micropipettes and graduated pipettes to prepare the testing solutions. </li>
 <li> Six graduated cylinders (e.g. 50 ml cylinders) for transferring testing solutions from the flasks to the testing containers. </li>
</ol>
3. Cyst Activation 
3.1. Petri dishes containing cysts + pouring water
To generate test organisms, 20 mg of cysts are placed into a Petri dish containing 12 ml of dilution water 48 hours before testing.
3.2. Incubation in the light and in the dark
The Petri dish is maintained at 25 (&plusmn;2) &deg;C and at a 4000 (&plusmn;1000) lux (lumens per square meter) light intensity for one hour. 
3.3. Replacing the medium using the microscope
After 24 hours, the hatching larvae are transferred into a new Petri dish filled with artificial water. The transfer is performed using the microscope, i.e. using a light source so that the hatching, phototropic larvae migrate towards the light beam. A glass Pasteur pipette is used to make the transfer, ensuring that only the nascent larvae are transferred and not the cysts or the larvae still in membranes.
3.4. Incubation in the dark
The dish containing the larvae is placed in a dark thermostatic chamber for 24 hours at 25 (&plusmn;2) &deg;C.
4. Preparation of Testing Solutions 
4.1. Weighing the substance + transfer to the flask
It is recommended that the standard solution for the testing substance is prepared by dissolving 0.5 g of the substance in a 500 ml flask. The flask is filled up with de-ionized or distilled water and the solution is stirred until the testing substance is completely dissolved. Solutions have to be prepared at the time of use unless it is not known whether the substance is stable in solution. In that case, the standard solution may be prepared up to two days before testing.
4.2. Preparing the negative control, adding dilution water and algae with the 10 ml pipette
The negative control is prepared in a flask (e.g. 200 ml volume) by adding an aliquot of the algal suspension to the dilution water so that a density of 105 cells/ml is obtained.
4.3. Preparing the testing solutions: adding dilution water, algae and standard solution
It is suggested that testing solutions are prepared in five 200 ml flasks by adding the standard solution to the dilution water in the quantities specified so that the desired concentrations for testing is obtained. To feed the organisms, aliquots of a Dunaliella tertiolecta microalgae suspension may be used and added to reach a density of 105 cells/ml when testing solutions are prepared. The following order of addition is recommended: dilution water, algal suspension and standard solution. After the preparation of testing solutions, they have to be used in the assay as soon as possible.
4.4. Transferring solutions from the flask to the testing containers, then to the Petri dishes
Equal quantities (50 ml) of the testing solutions are introduced into the testing containers utilizing the graduated cylinder. Three replicas for each concentration are made. For each series of tests, a control container is prepared with an amount of dilution water equal to the volume of the test solutions. Equal volumes (about 12 ml) of the test solutions are introduced into the Petri dishes. For each series of tests, a control Petri dish is used.
5. Preparation for Testing 
5.1. Nauplii at the II-III stage
Forty-eight hours after cyst activation, the nauplii reach the II-III larval stage and are then usable for testing.
5.2. Nauplii transfer into the Petri dishes using the microscope
The Petri dish used for cyst activation is taken out of the thermostatic chamber. A small quantity of larvae are transferred into the Petri dishes containing the control and the testing solutions. This task is to be performed with the microscope utilizing a pipette of sufficiently wide diameter so that the organisms do not get damaged. In this phase of the test, the transfer may be performed with a glass Pasteur pipette (with tip rounded by flame) and a laterally positioned light source that encourages the Artemia to aggregate.
5.3. Nauplii transfer from the Petri dishes into the test containers with the microscope
Ten larvae are transferred from the testing solutions Petri dishes to the testing containers. Also this operation has to be performed by utilizing the Pasteur pipette and microscope. It is recommended that a volume not exceeding 1 mL is transferred during the passage of larvae not to affect the overall volume of test system.
5.4. Full containers covered with parafilm
The testing beakers are covered with parafilm (leaving a gap for air passage), and maintained at a temperature of 25 (&plusmn;2) &deg;C for the whole duration of the test and at an illumination of 900 (&plusmn;100) lx with a photoperiod ratio of 14 hours of light to 10 hours of darkness. 
6. Replacing the Medium and Food Supplement 
6.1. Survival rate control
Two days after the test has started, and then after 5, 7, 9, and 12 days, the Artemia is observed under the microscope to verify the survival rate and to replace the medium and the food supplement. 
Number of live organisms is counted in each test container. After observation under the microscope and slight mechanical stimulation (e.g. touching the larvae with a glass Pasteur pipette) organisms that do not show some movement for about 10 seconds should be considered dead. 
6.2. Preparation of a test solution showing algae sampling with a 10 ml pipette and sampling of the substance to be tested with a micropipette. Transferring solutions from the flask to the test containers
While testing, the test solutions must be periodically replaced, and prepared in the same day as they are to be used. Test solutions are made from the standard solution previously prepared. 
6.3. Artemia transfer, utilizing a cut Pasteur pipette, from the old test containers to the three new containers
The transfer of Artemia is performed by using a pipette with a sufficiently wide diameter so as not to damage the organisms. During this phase, a 3 ml plastic Pasteur pipette (to be cut) may be used.
6.4. Larvae in a beaker containing either live or dead Artemia
After 14 days of exposure at the end of testing, the number of surviving larvae is counted and recorded on the worksheet. LC50 and/or LC20, NOEC and LOEC at 14 days, are calculated and recorded, as well as confidence limits at 95 % where appropriate, and calculation methods.
7. Representative Results
At the end of the test (14 days), calculate the percentage of mortality at each concentration, according to the following formula: 
(Ms/N) * 100
Where:
Ms is average number of died individuals in the sample analysed
N is total number of exposed individuals 
Table 1 reports an example of data of a test (14d) with Artemia exposed to Sodium Dodecyl Sulfate.
<table border="1">
<tbody>
 <tr>
 <td>Concentrations (mg/l)</td>
 <td colspan="3">Number of died individuals in each replica </td>
 <td>Mortality (%)</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>I</td>
 <td>II</td>
 <td>III</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Negative control</td>
 <td>0</td>
 <td>2</td>
 <td>1</td>
 <td>10</td>
 </tr>
 <tr>
 <td>1.56</td>
 <td>1</td>
 <td>2</td>
 <td>0</td>
 <td>10</td>
 </tr>
 <tr>
 <td>3.12</td>
 <td>2</td>
 <td>0</td>
 <td>2</td>
 <td>13</td>
 </tr>
 <tr>
 <td>6.25</td>
 <td>3</td>
 <td>2</td>
 <td>3</td>
 <td>27</td>
 </tr>
 <tr>
 <td>12.5</td>
 <td>8</td>
 <td>9</td>
 <td>10</td>
 <td>90</td>
 </tr>
 <tr>
 <td>25.0</td>
 <td>10</td>
 <td>10</td>
 <td>10</td>
 <td>100</td>
 </tr>
 <tr>
 <td>50.0</td>
 <td>10</td>
 <td>10</td>
 <td>10</td>
 <td>100</td>
 </tr>
 </tbody>
</table>
Moreover, determine the LC50 value by a graphical Gaussian logarithmic plot or using appropriate statistical methods (for example Spearman-Karber or Probit Method). The calculation of LC50 for the data of table 1 is 8.18 (7.15-9.37) mg/l. If the maximum concentration tested caused a lower mortality rate of 50%, you should not proceed to the calculation of the LC50 value which would then be unreliable or even non-computable. In this case, however, would be appropriate to repeat the test by extending the range of concentrations tested. Alternatively, the LC50 value is more correctly expressed as greater than the highest concentration tested, possibly indicating the percentage of mortality at the highest concentration tested and the highest concentration corresponding to a no effect.
Results are considered valid if at the end of testing the following conditions have been met:
<ol class="lalpha">
 <li> the control's average mortality rate is &le; 20 %;
 </li>
 <li> where using Sodium Dodecyl Sulfate, the LC50 at 14 days is included in the 8.0 (&plusmn;5) mg/l interval.
 </li>
</ol>
If the above conditions have not been met, all data obtained with the same batch of organisms should be considered invalid and testing repeated.

<ol>
	<li>Togulga, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Short-Term+Toxicity+of+Two+Toxicants+to+Artemia+Nauplii.">The Short-Term Toxicity of Two Toxicants to Artemia Nauplii.</a> Tr. J. of Zoology. 22, 259-266 (1998).</li><li>Persoone, G., Wells, P. G., Sorgeloos, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Artemia+in+aquatic+toxicology:+a+review.">Artemia in aquatic toxicology: a review.</a> Artemia Research and its Applications. Morphology, Genetics, Strain characterization Toxicology. , 259-275 (1987).</li><li>Nunes, B. S., Carvalho, F. D., Guilhermino, L. M., Stappen, G. V. a. n. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+the+genus+Artemia+in+ecotoxicity+testing.">Use of the genus Artemia in ecotoxicity testing.</a> Envir. Poll. 144, 453-462 (2006).</li><li>APAT IRSA-CNR. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Metodi analitici per le acque Manuali e Linee guida 29/2003. Terzo Metodo, 8060 (2003).</li><li>Sorgellos, P., Wielen, C. R. e. m. i. c. h. e. -. V. a. n. D. e. r., Persoone, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+Artemia+nauplii+for+toxicity+tests+â€“+a+critical+analysis.">The use of Artemia nauplii for toxicity tests â€“ a critical analysis.</a> Ecotox. Env. Saf. 2, 249-255 (1978).</li><li>Savorelli, F., Palazzi, D., Gorbi, G., Invidia, M., Sei, S., Magaletti, E., Manfra, L., Gelli, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Set+up+of+a+standard+methodology+for+14-day+bioassay+on+Artemia+franciscana+and+A.+parthenogenetica.">Set up of a standard methodology for 14-day bioassay on Artemia franciscana and A. parthenogenetica.</a> Biol. Amb. 21, 27-36 (2007).</li><li>Brix, K. V., Cardwell, R. D., Adams, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+toxicity+of+arsenic+to+the+Great+Salt+Lake+brine+shrimp,+Artemia+franciscana.">Chronic toxicity of arsenic to the Great Salt Lake brine shrimp, Artemia franciscana.</a> Ecotox. Environ. Saf. 54, 169-175 (2003).</li><li>Brix, K. V., Deforest, D. K., Cardwell, R. D., Adams, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Derivation+of+a+chronic+site-specific+water+quality+standard+for+selenium+in+the+Great+Salt+Lake,+Utah,+USA.">Derivation of a chronic site-specific water quality standard for selenium in the Great Salt Lake, Utah, USA.</a> Environ. Toxicol. Chem. 23, 606-612 (2004).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Qualit&#225;+dell'acqua+-+Determinazione+della+tossicit&#225;+letale+a+14+giorni+con+Artemia+franciscana+(Crustacea:+Anostraca).">Qualit&#225; dell'acqua - Determinazione della tossicit&#225; letale a 14 giorni con Artemia franciscana (Crustacea: Anostraca).</a> Commissione UNICHIM "Qualit&#225; dell'acqua" Gruppo di Lavoro "Metodi Biologici" Sottogruppo "Acque salate/salmastre e sedimenti". , (2010).</li><li>Varo, I., Navarro, J. C., Amat, F., Guilhermino, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterisation+of+cholinesterases+and+evaluation+of+the+inhibitory+potential+of+chlorpyrifos+and+dichlorvos+to+Artemia+salina+and+Artemia+parthenogenetica.">Characterisation of cholinesterases and evaluation of the inhibitory potential of chlorpyrifos and dichlorvos to Artemia salina and Artemia parthenogenetica.</a> Chem. 48, 563-569 (2002).</li></ol>

No conflicts of interest declared.

Artemia is one of the most valuable test organisms available for ecotoxicity testing and research done so far allows us to state that it is possible to sustain several options related to Artemia use in Toxicology and Ecotoxicology. The characteristics that turn this organism into a suitable species for bioassays are: a wide geographic distribution, high adaptability to adverse environmental conditions and varied nutrients, relatively simple laboratory culture and maintenance, resistance to manipulation,...

Produced by Loredana Manfra of Institute for environmental protection and research (ISPRA-Department of Environmental Quality Monitoring) and Federica Savorelli of Regional Agency for Environmental Protection (Emilia-Romagna, Italy).
Reference Members for the sub-working Group on the Artemia crustacean within the Biological Methods Group &#x2013; Saltwater/Brackish Water and Sediment of the UNICHIM Commission on Water Quality (the institute for technical standardization operating in the chemical sector).
Videography and editing by Marco Pisapia of ISPRA (Scientific Advisory and Documentation Services). 
Production Coordination by Anna Maria Cicero and Erika Magaletti of ISPRA (Department of Environmental Quality Monitoring).
We would like to thank the Regional Agency for Environmental Protection of Emilia-Romagna, Ferrara Branch and the Biological Methods Group &#x2013; Saltwater /Brackish Water and Sediment of the UNICHIM Commission on Water Quality for their cooperation; Rossella Boscolo and Massimo Gabellini of ISPRA (Department of Prevention and Mitigation of Impacts) for their financial support; and the London School of Languages in Rome for their linguistic support.
<table border="1">
<tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Product number</td>
 </tr>
 <tr>
 <td>Sodium Dodecyl Sulfate </td>
 <td>Sigma-Aldrich </td>
 <td>L4390 </td>
 </tr>
 </tbody>
</table>
Appendix
The algal media has to be prepared using analytical quality reagents. 
A liter of nutrient solution is prepared by dissolving 300 g of NaNO3, 30 g of KH2PO4 and 20 g of NH4Cl in distilled water. 
Four initial solutions (A, B, C, D) of minerals are prepared as follows:
<ul>
 <li> A liter of solution "A" containing 30 g of ZnSO4 H2O, 25 g of CuSO4 5 H2O, 30 g of CoSO4 7 H2O and 20 g of MnSO4 H2O in distilled water;
 </li>
 <li> A liter of solution "B" containing 50 g of FeCl3 &bull;6 H2O in distilled water; 
 </li>
 <li> A liter of solution "C" containing 25 g of Na2MoO4 &bull;2 H2O in distilled water;
 </li>
 <li> A liter of solution "D" containing 50 g di Na2EDTA &bull;2 H2O in distilled water.
 </li>
</ul>
A liter of a final solution of minerals (solution "E") is obtained adding aliquots of the above-mentioned solutions to distilled water. The following order of addition is recommended: 100 ml of solution "D", 10 ml of solution "C", 10 ml of solution "B" and 10 ml of solution "A".
Three initial solutions (F, G, H) of vitamins are prepared as follows:
<ul>
 <li> A liter of solution "F" containing 100 mg of biotin in distilled water;
 </li>
 <li> A liter of solution "G" containing 10 g of thiamine in distilled water;
 </li>
 <li> A liter of solution "H" containing 100 mg of cobalamin in distilled water.
 </li>
</ul>
A liter of a final solution of vitamins (solution "I") is obtained adding 10 ml of the three initial solutions to distilled water.
A liter of the algal media is prepared by dissolving 1 ml of the nutrient solution, 1 ml of the mineral final solution (solution "E") and 1 ml of the vitamin final solution (solution "I") in seawater. This latter may be obtained by dissolving recognized analytical quality reagents or a commercially available formulation in distilled or de-ionized water. Salt blends for creating the ideal saltwater are commercially available, e.g. the Instant Ocean mixtures. It is recommended the using of a seawater with salinity of 30 (&plusmn;2) PSU and temperature of 20 (&plusmn;1) &deg;C.

long-term lethal toxicity test with the crustacean artemia franciscana

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish water and sediment. The choice of biological indicators must be based on reliable scientific knowledge and, possibly, on the availability of standardized procedures. In this article, we present a standardized protocol that used the marine crustacean Artemia to evaluate the toxicity of chemicals and/or of marine environmental matrices. Scientists propose that the brine shrimp (Artemia) is a suitable candidate for the development of a standard bioassay for worldwide utilization. A number of papers have been published on the toxic effects of various chemicals and toxicants on brine shrimp (Artemia). The major advantage of this crustacean for toxicity studies is the overall availability of the dry cysts; these can be immediately used in testing and difficult cultivation is not demanded1,2. Cyst-based toxicity assays are cheap, continuously available, simple and reliable and are thus an important answer to routine needs of toxicity screening, for industrial monitoring requirements or for regulatory purposes3. The proposed method involves the mortality as an endpoint. The numbers of survivors were counted and percentage of deaths were calculated. Larvae were considered dead if they did not exhibit any internal or external movement during several seconds of observation4. This procedure was standardized testing a reference substance (Sodium Dodecyl Sulfate); some results are reported in this work. This article accompanies a video that describes the performance of procedural toxicity testing, showing all the steps related to the protocol.

Watch this Scientific Journal Video about Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana at JoVE.com

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

This study concerns the development and standardization of a valuable methodological protocol to determine long-term (14 days) lethal toxicity exerted by chemical substances, industrial wastewater or sewage and liquid environmental samples on the saltwater crustacean, Artemia franciscana.

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish ...

long-term-lethal-toxicity-test-with-the-crustacean-artemia-franciscana

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19.5K Views.  Institute for Environmental Protection and Research. This study concerns the development and standardization of a valuable methodological protocol to determine long-term (14 days) lethal toxicity exerted by chemical substances, industrial wastewater or sewage and liquid environmental samples on the saltwater crustacean, Artemia franciscana.

Video: Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

Long-term Imaging Mammalian Cells using Wide-Field Microscopy

Actually what I'm gonna do, mount it in here first and then swap out You. So now I'm gonna aspirate try and be as gentle as possible.Okay. A few seconds just holding.Okay.Alright.So the important features of our lifestyle imaging grid are that, so it's an inverted scope, which means the objective, the samples here in the objective is underneath. So the objective is pointing up and the whole scope, or practically the whole, practically the whole scope, including the objectives and the sample are at 37 degrees. So everything in this plastic box is at 37 degrees.And right on top of the stage we have an air and CO2 mix. That is 5%CO2. So the idea is that the sample is actually in a cell, it's like a cell incubator.And so we have, this is our mixer for the air and the CO2 and, and it gets humidified and heated and then piped onto the, the sample on the stage.Okay. And today we're just gonna be doing fluorescence one channel GFP fluorescence. This is the hose that the CO2 and 5%CO2 goes in and it just fits into the stage holder.That's important. And then we can close these, take these down, and then we go to our first position. And what I'm gonna do is go to the first field, which is my control sample, and I'll pick positions based on what I see on the screen.So I just make sure I actually look into the microscope in each new, well, because the focal plane for each of the wells is different, the, the dish is not perfectly flat on the bottom. So you will need to make fine adjustments when you move from well to the well. So right now I'm on the first, well, what I'll do is I'll focus, so I just hit, I just told, I just turned on the light, the transmitted light for the microscope.And I have make sure all the light is going to the eye pieces. And so I just look in, make sure I get a nice clean field. And then what I'll do is turn that light off, turn the fluorescence light on, make sure the cells that I'm looking at look healthy and they're not apoptotic.And, and I turn that off. And what I do next is send, so now I'm gonna start picking positions, which means I have to te send the light path to the camera. This is the camera right here.So I tell the microscope to send all the camera, they don't see anything in the eye pieces. What I'll do is in the software controlling the microscope, I'll choose the right filter set, which for our purposes is psy. And then set an exposure time.And I know these cells very well and I know that they take a 0.2 second exposure time. So I tell, that's what I tell the software to do. And what I'll do is hit focus.This is, and it's basically just alive. The, A light will turn on and you can, oh, I don't know if you can see cells, but the reason this is a good field is because there's, all the cells are, even in terms of their fluorescence, There are no apoptotic cells. Apoptotic cells tend to be very bright and there's no dirt or schmutz in the field.And you can see that they're actually, this is a metaphase cell and that's a metaphase cell, that's a cell that's probably just exited mitosis. And I can just tell that because it looks like those are, those two sets of chromatin are very nicely oriented to each other. Like they just exited an face.So that's a good field. So, and now that I know that the exposure time and the filter set is correct, and this is gonna be software specific, I'll go and I'll take positions and now the software has memorized the X, Y, Z position of that field. And then when I do pick another field, it's, and I'll, so when I'm picking, when I'm picking new fields, but in the same, well, I'll use the, I'll move the i'll, I'll turn the shutter, I'll open the shutter and I'll look to see what I see on the screen and I'll pick fields based on that versus looking in the field.So I don't see any of my tonics in there. But again, even nicely centered, these cells will go through my PS I continue, here's a good example. Here's a good example of what I would avoid that, see how bright that is?That's very bright. That's probably remnants of an apoptotic cell. And the reason I would avoid that is because since it's so bright, when the software auto focuses, it will autofocus just on that and it won't be in the same plane as all these other cells.And so I won't get any information from that field. That looks nice. There's a, this is a pro metaphase right here, That's A metaphase.So, and I'm gonna take three positions per well. So that was three positions and now I'm going to go to my next Well, so you can, so what I will probably do is take images every 10 minutes. So it's a compromise.Lifestyle imaging is always a compromise, right? You have to image, you have to keep the cells healthy, especially if you're interested in mitosis. So you have to minimize their exposure to the fluorescent line.And, but you wanna get information. So you have to take images, you have to, so you have to compromise, you have to make a choice about what you want to capture, what kind of information you wanna capture. For our purposes, mitosis tends to take about an hour.So if we're taking 10 minute intervals, that's pretty good. We see prophase, we see metaphase, we see anaphase, and we can, we can see when there are, there are treatments or drugs or siRNAs that affect those specific cell cycle stages. So for our purposes, that's enough, but we always want more.We always want more positions or finer time and interval or something like that. So it's a compromise. So, and then I'll hit start.And so our software, it does an autofocusing pass on the first pass. So once I hit start, it should start auto autofocusing and you'll see the shutter open, open and close between auto focusing. So I'll hit start.So that's the shutter opening and closing. The Z motor is engaging and it's actually trying to find the plane with the most contrast, which will be the most in focus. You can actually see on the screen going in and out of finding the best plane of focus that might be worrisome.So what it does is it, it drives the Z motor to find the best, the most in focus plane focus, well the most in image. And then it acquires picture there and then it learns that Z position. And then we'll use that Z position for the next 12 passes.And then we'll do the auto focusing pass again. What Kind of experiments do they do? Okay, so our lab is mostly interested in understanding what regulates mitic progression, or at least I'm most interested in understanding that.So with live cell imaging and especially our multi position, long-term imaging scope, you can take, you can film cells for two or three days entering anding mitosis, and you can image multiple rounds of mitosis so you can understand. And then you can put drugs or RNAs or any kind of perturbations on the cells that you would like to study. So you can understand how, how those drugs or, or genes are, are important in regulating mitotic progression.Because it's not fixed cell work and live cell work, you can understand what, when you need to have a perturbation to affect mitosis or what stage of mitosis is affected. So it's really powerful in that respect. What, what are the main technical elements of this experiments?What are the possible problems? Well, the biggest problem besides getting the rig set up, getting the rig set up is not trivial. A lot of people have the rig, but just making sure it works well for your application.The financial element is not, you know, is not small. So there's the setting up of the apparatus and then, and then once you gather the data, there's a real data analysis issue. It takes these movie, if, so, if you're gathering 40 movies each with 30 cells, so you have 40 in 40 fields that you just filmed and each of those fields has 30 cells, then you have to sit down with each one of those movies and look at each one of those cells in that movie.So it takes a really long time to do data analysis. And that's, so it takes a week to analyze data that it took you two days to collect. So there's a, that's a very big obstacle in terms of gathering lots of data.So it's not, while it's high throughput data collection, it's not high throughput analysis. So what would you make to make this experiment really high throughput? So our lab is in the process of developing a program that does the data analysis automatically.And it's a pretty decent program. It's not perfect. The human eye is always better, but it's getting there.So What are the interesting application of this experiment? Maybe other fields you can mention? Besides cell division?You mean for the, oh, for the live cell imaging? Yes.Well, you can do cell motility assays. You can, depending on the, the microscopy that's involved, you can, you know, look at how cells, you could look at tumor development if you really wanted to.But there, you, you're, if as long as you have the right imaging set up, you can, you're pretty much not limited in terms of anything. Our scope happens to be just a wide field and we use 20 by 20 x objective. So it's a pretty low resolution set up, but that's sufficient for what we need to see.So there are, but there definitely aren't, you wouldn't be limited. Anything that has any dynamic kind of experiment, which is biology, it would be, would really benefit from lifestyle imaging. So calcium imaging, although you couldn't do it on our apparatus, that's a live cell.You definitely need live cell imaging for that. Like I said, patient assays, motility assays, those kinds of things would really benefit from live cell imaging. Would you like to tell a few words about your previous experience?How long do you work on these techniques? What do you you work on before? Wow.So in my PhD I didn't work.I did a lot of confocal microscopy, but no lifestyle imaging. And so during my postdoc I was, I helped Dr.King get this apparatus that, that we just used to set up these fragment, get that up and running. And, and so during my postdoc, I had a lot of experience with both widefield with TimeLapse imaging on our scope, which is the widefield and also TimeLapse imaging with confocal microscopy.And I would say that it's really powerful. You get a lot of information, but it's not easy to get working perfectly. So it really requires a lot of attention and a lot of attention to details at every little step of the process.And, and then you still have problems. So it's a tough application, but you can really get, if, if it's what you need to have to, to study your problem, your scientific problem, then it, it's invaluable in that respect, but it's tough. So find someone who knows what they're doing and use them.That's my advice. So good.

Długoterminowe Imaging komórkach ssaków przy dużym polu Mikroskopia

Long-term Culture of Human Breast Cancer Specimens and Their Analysis Using Optical Projection Tomography

Breast cancer is a leading cause of mortality in the Western world. It is well established that the spread of breast cancer, first locally and later distally, is a major factor in patient prognosis. Experimental systems of breast cancer rely on cell lines usually derived from primary tumours or pleural effusions. Two major obstacles hinder this research: (i) some known sub-types of breast cancers (notably poor prognosis luminal B tumours) are not represented within current line collections; (ii) the influence of the tumour microenvironment is not usually taken into account.
We demonstrate a technique to culture primary breast cancer specimens of all sub-types. This is achieved by using three-dimensional (3D) culture system in which small pieces of tumour are embedded in soft rat collagen I cushions. Within 2-3 weeks, the tumour cells spread into the collagen and form various structures similar to those observed in human tumours1. Viable adipocytes, epithelial cells and fibroblasts within the original core were evident on histology. Malignant epithelial cells with squamoid morphology were demonstrated invading into the surrounding collagen. Nuclear pleomorphism was evident within these cells, along with mitotic figures and apoptotic bodies.
We have employed Optical Projection Tomography (OPT), a 3D imaging technology, in order to quantify the extent of tumour spread in culture. We have used OPT to measure the bulk volume of the tumour culture, a parameter routinely measured during the neo-adjuvant treatment of breast cancer patients to assess response to drug therapy. 
Here, we present an opportunity to culture human breast tumours without sub-type bias and quantify the spread of those ex vivo. This method could be used in the future to quantify drug sensitivity in original tumour. This may provide a more predictive model than currently used cell lines.

We have developed a collagen-based in vitro assay which promotes proliferation and invasion from samples of all breast cancer subtypes. Optical Projection Tomography, a three dimensional microscopy technique was utilised to visualise and quantify tumour expansion. This assay may be used to quantify drug response of individual tumour samples.

Breast cancer is a leading cause of mortality in the Western world. It is well established that the spread of breast cancer, first locally and later ...

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue patterning. During animal development, key cell proliferation and patterning events occur very quickly. For instance, in Caenorhabditis elegans all cell divisions required for the larval body plan are completed within six hours after fertilization, with seven mitotic cycles1; the sixteen or more mitoses of Drosophila embryogenesis occur in less than 24 hr2. In contrast, cell divisions during plant development are slow, typically on the order of a day 3,4,5 . This imposes a unique challenge and a need for long-term live imaging for documenting dynamic behaviors of cell division and differentiation events during plant organogenesis. Arabidopsis epidermis is an excellent model system for investigating signaling, cell fate, and development in plants. In the cotyledon, this tissue consists of air- and water-resistant pavement cells interspersed with evenly distributed stomata, valves that open and close to control gas exchange and water loss. Proper spacing of these stomata is critical to their function, and their development follows a sequence of asymmetric division and cell differentiation steps to produce the organized epidermis (Fig. 1).
This protocol allows observation of cells and proteins in the epidermis over several days of development. This time frame enables precise documentation of stem-cell divisions and differentiation of epidermal cells, including stomata and epidermal pavement cells. Fluorescent proteins can be fused to proteins of interest to assess their dynamics during cell division and differentiation processes. This technique allows us to understand the localization of a novel protein, POLAR6, during the proliferation stage of stomatal-lineage cells in the Arabidopsis cotyledon epidermis, where it is expressed in cells preceding asymmetric division events and moves to a characteristic area of the cell cortex shortly before division occurs. Images can be registered and streamlined video easily produced using public domain software to visualize dynamic protein localization and cell types as they change over time.

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

We describe a protocol using chamber slides and media to immobilize plant cotyledons for confocal imaging of the epidermis over several days of development, documenting stomatal differentiation. Fluorophore-tagged proteins can be tracked dynamically by expression and subcellular localization, increasing understanding of their possible roles during cell division and cell-type differentiation.

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue ...

Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

The amphipod Parhyale hawaiensis is a small crustacean found in intertidal marine habitats worldwide. Over the past decade, Parhyale has emerged as a promising model organism for laboratory studies of development, providing a useful outgroup comparison to the well studied arthropod model organism Drosophila melanogaster. In contrast to the syncytial cleavages of Drosophila, the early cleavages of Parhyale are holoblastic. Fate mapping using tracer dyes injected into early blastomeres have shown that all three germ layers and the germ line are established by the eight-cell stage. At this stage, three blastomeres are fated to give rise to the ectoderm, three are fated to give rise to the mesoderm, and the remaining two blastomeres are the precursors of the endoderm and germ line respectively. However, blastomere ablation experiments have shown that Parhyale embryos also possess significant regulatory capabilities, such that the fates of blastomeres ablated at the eight-cell stage can be taken over by the descendants of some of the remaining blastomeres. Blastomere ablation has previously been described by one of two methods: injection and subsequent activation of phototoxic dyes or manual ablation. However, photoablation kills blastomeres but does not remove the dead cell body from the embryo. Complete physical removal of specific blastomeres may therefore be a preferred method of ablation for some applications. Here we present a protocol for manual removal of single blastomeres from the eight-cell stage of Parhyale embryos, illustrating the instruments and manual procedures necessary for complete removal of the cell body while keeping the remaining blastomeres alive and intact. This protocol can be applied to any Parhyale cell at the eight-cell stage, or to blastomeres of other early cleavage stages. In addition, in principle this protocol could be applicable to early cleavage stage embryos of other holoblastically cleaving marine invertebrates.

Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

The amphipod Parhyale hawaiensis is a promising model organism for studies of crustacean embryology and comparative arthropod development and evolution. This protocol describes a method for manual removal of single blastomeres from early cleavage stage embryos of Parhyale.

The amphipod Parhyale hawaiensis is a small crustacean found in intertidal marine habitats worldwide. Over the past decade, Parhyale has emerged as a ...

Multilayer Mounting for Long-term Light Sheet Microscopy of Zebrafish

Light sheet microscopy is the ideal imaging technique to study zebrafish embryonic development. Due to minimal photo-toxicity and bleaching, it is particularly suited for long-term time-lapse imaging over many hours up to several days. However, an appropriate sample mounting strategy is needed that offers both confinement and normal development of the sample. Multilayer mounting, a new embedding technique using low-concentration agarose in optically clear tubes, now overcomes this limitation and unleashes the full potential of light sheet microscopy for real-time developmental biology.

The development of zebrafish can be followed over days with light sheet microscopy when embryos are embedded in optically clear polymer tubes with low-concentration agarose.

Light sheet microscopy is the ideal imaging technique to study zebrafish embryonic development. Due to minimal photo-toxicity and bleaching, it is ...

LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging

Live imaging is a valuable approach for investigating cell biology questions. The Drosophila larva is particularly suited for in vivo live imaging because the larval body wall and most internal organs are transparent. However, continuous live imaging of intact Drosophila larvae for longer than 30 min has been challenging because it is difficult to noninvasively immobilizeimmobilizing larvae for a long time. Here we present a larval mounting method called LarvaSPA that allows for continuous imaging of live Drosophila larvae with high temporal and spatial resolution for longer than 10 hours. This method involves partially attaching larvae to the coverslip using a UV-reactive glue and additionally restraining larval movement using a polydimethylsiloxane (PDMS) block. This method is compatible with larvae at developmental stages from second instar to wandering third instar. We demonstrate applications of this method in studying dynamic processes of Drosophila somatosensory neurons, including dendrite growth and injury-induced dendrite degeneration. This method can also be applied to study many other cellular processes that happen near the larval body wall.

LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging

This protocol describes a method for mounting Drosophila larvae to achieve longer than 10 h of uninterrupted time-lapse imaging in intact live animals. This method can be used to image many biological processes close to the larval body wall.

Live imaging is a valuable approach for investigating cell biology questions. The Drosophila larva is particularly suited for in vivo live imaging because ...

Isolation, Transfection, and Long-Term Culture of Adult Mouse and Rat Cardiomyocytes

Ex vivo culture of the adult mammalian cardiomyocytes (CMs) presents the most relevant experimental system for the in vitro study of cardiac biology. Adult mammalian CMs are terminally differentiated cells with minimal proliferative capacity. The post-mitotic state of adult CMs not only restricts cardiomyocyte cell cycle progression but also limits the efficient culture of CMs. Moreover, the long-term culture of adult CMs is necessary for many studies, such as CM proliferation and analysis of gene expression.
The mouse and the rat are the two most preferred laboratory animals to be used for cardiomyocyte isolation. While the long-term culture of rat CMs is possible, adult mouse CMs are susceptible to death and cannot be cultured more than five days under normal conditions. Therefore, there is a critical need to optimize the cell isolation and long-term culture protocol for adult murine CMs. With this modified protocol, it is possible to successfully isolate and culture both adult mouse and rat CMs for more than 20 days. Moreover, the siRNA transfection efficiency of isolated CM is significantly increased compared to previous reports. For adult mouse CM isolation, the Langendorff perfusion method is utilized with an optimal enzyme solution and sufficient time for complete extracellular matrix dissociation. In order to obtain pure ventricular CMs, both atria were dissected and discarded before proceeding with the disassociation and plating. Cells were dispersed on a laminin coated plate, which allowed for efficient and rapid attachment. CMs were allowed to settle for 4-6 h before siRNA transfection. Culture media was refreshed every 24 h for 20 days, and subsequently, CMs were fixed and stained for cardiac-specific markers such as Troponin and markers of cell cycle such as KI67.

Here, we present a protocol for the isolation, transfection, and long-term culture of adult mouse and rat cardiomyocytes.

Ex vivo culture of the adult mammalian cardiomyocytes (CMs) presents the most relevant experimental system for the in vitro study of cardiac biology. ...

Application of Passive Head Motion to Generate Defined Accelerations at the Heads of Rodents

Exercise is widely recognized as effective for various diseases and physical disorders, including those related to brain dysfunction. However, molecular mechanisms behind the beneficial effects of exercise are poorly understood. Many physical workouts, particularly those classified as aerobic exercises such as jogging and walking, produce impulsive forces at the time of foot contact with the ground. Therefore, it was speculated that mechanical impact might be implicated in how exercise contributes to organismal homeostasis. For testing this hypothesis on the brain, a custom-designed ''passive head motion'' (hereafter referred to as PHM) system was developed that can generate vertical accelerations with controlled and defined magnitudes and modes and reproduce mechanical stimulation that might be applied to the heads of rodents during treadmill running at moderate velocities, a typical intervention to test the effects of exercise in animals. By using this system, it was demonstrated that PHM recapitulates the serotonin (5-hydroxytryptamine, hereafter referred to as 5-HT) receptor subtype 2A (5-HT2A) signaling in the prefrontal cortex (PFC) neurons of mice. This work provides detailed protocols for applying PHM and measuring its resultant mechanical accelerations at rodents' heads.

The present protocol describes a custom-designed ''passive head motion'' system, which reproduces mechanical accelerations at rodents' heads generated during their treadmill running at moderate velocities. It allows dissecting mechanical factors/elements from the beneficial effects of physical exercise.

Exercise is widely recognized as effective for various diseases and physical disorders, including those related to brain dysfunction. However, molecular ...

Collection and Long-Term Maintenance of Leaf-Cutting Ants (Atta) in Laboratory Conditions

Ants are one of the most biodiverse groups of animals on the planet and inhabit different environments. The maintenance of ant colonies in controlled environments enables an enriched comprehension of their biology that can contribute to applied research. This practice is usually employed in population control studies of species that cause economic loss, such as Atta ants. To cultivate their mutualistic fungus, these leaf-cutting ants collect leaves and for this are considered agricultural pests widely distributed throughout the American continent. They are highly socially organized and inhabit elaborated underground nests composed of a variety of chambers. Their maintenance in a controlled environment depends on a daily routine of several procedures and frequent care that are described here. It starts with the collection of queens during the reproductive season (i.e., nuptial flight), which are then individually transferred to plastic containers. Due to the high mortality rate of queens, a second collection can be carried out about 6 months after the nuptial flight, when incipient nests with developed fungus wad are excavated, hand-picked, and placed in plastic containers. In the laboratory, leaves are daily provided to established colonies, and ant-produced waste is weekly removed along with remaining dry plant material. As the fungus garden keeps growing, colonies are transferred to different types of containers according to the experimental purpose. Leaf-cutting ant colonies are placed in interconnected containers, representing the organizational system with functional chambers built by those insects in nature. This setup is ideal to monitor factors such as waste amount, fungus garden health, and the behavior of workers and queen. Facilitated data collection and more detailed observations are considered the greatest advantage of keeping ant colonies in controlled conditions.

Collection and Long-Term Maintenance of Leaf-Cutting Ants Atta in Laboratory Conditions

Here, a protocol is described to successfully collect and maintain healthy Atta (Hymenoptera: Formicidae) ant colonies in laboratory conditions. Additionally, different nest types and configurations are detailed together with possible experimental procedures.

Ants are one of the most biodiverse groups of animals on the planet and inhabit different environments. The maintenance of ant colonies in controlled ...

Long-Term Culture and Monitoring of Isolated Caenorhabditis elegans on Solid Media in Multi-Well Devices

The nematode Caenorhabditis elegans is among the most common model systems used in aging research owing to its simple and inexpensive culture techniques, rapid reproduction cycle (~3 days), short lifespan (~3 weeks), and numerous available tools for genetic manipulation and molecular analysis. The most common approach for conducting aging studies in C. elegans, including survival analysis, involves culturing populations of tens to hundreds of animals together on solid nematode growth media (NGM) in Petri plates. While this approach gathers data on a population of animals, most protocols do not track individual animals over time. Presented here is an optimized protocol for the long-term culturing of individual animals on microfabricated polydimethylsiloxane (PDMS) devices called WorMotels. Each device allows up to 240 animals to be cultured in small wells containing NGM, with each well isolated by a copper sulfate-containing moat that prevents the animals from fleeing. Building on the original WorMotel description, this paper provides a detailed protocol for molding, preparing, and populating each device, with descriptions of common technical complications and advice for troubleshooting. Within this protocol are techniques for the consistent loading of small-volume NGM, the consistent drying of both the NGM and bacterial food, options for delivering pharmacological interventions, instructions for and practical limitations to reusing PDMS devices, and tips for minimizing desiccation, even in low-humidity environments. This technique allows the longitudinal monitoring of various physiological parameters, including stimulated activity, unstimulated activity, body size, movement geometry, healthspan, and survival, in an environment similar to the standard technique for group culture on solid media in Petri plates. This method is compatible with high-throughput data collection when used in conjunction with automated microscopy and analysis software. Finally, the limitations of this technique are discussed, as well as a comparison of this approach to a recently developed method that uses microtrays to culture isolated nematodes on solid media.

Long-Term Culture and Monitoring of Isolated Caenorhabditis elegans on Solid Media in Multi-Well Devices

Presented here is an optimized protocol for culturing isolated individual nematodes on solid media in microfabricated multi-well devices. This approach allows individual animals to be monitored throughout their lives for a variety of phenotypes related to aging and health, including activity, body size and shape, movement geometry, and survival.

The nematode Caenorhabditis elegans is among the most common model systems used in aging research owing to its simple and inexpensive culture techniques, ...