# Presentation: 184930

## Techniques for the Development of New Insect Cell Lines

- Workshop at 2005 Annual Meeting – SIVB
- Saturday 4 June 3:00-4:30PM
- D. E. Lynn, C. Goodman, G. Caputo

- http://www.ars.usda.gov/SP2UserFiles/Place/12752100/InsectTechniques.pps

## Equipment

- Laminar flow hood (clean bench)
- Dissecting microscope
- Alcohol lamp/Alcohol jar
- 			or
- Clorox/water rinse
- Fine surgical instruments

- Plus the ‘usual’ tissue culture equipment and supplies (26-28°C incubator, pipettor, pipets, culture dishes, flasks, etc.)

**Notes:**
While most cell culture laboratories now use biosafety cabinets, a clean bench is preferable for primary culture work because it is easier to work with a dissecting microscope.
The fine instruments are available from surgical supply companies because they are used in medicine for fine operations (eye and vascular surgery, for example).
Dwight uses alcohol flaming to ‘sterilize’ his instruments while Cindy uses Clorox and distilled water rinses.
Use care when buying an incubator for insect cell culture research.  Most incubators designed for mammalian cell culture work are designed to work well at 35-38°C but have problems maintaining at temperature appropriate for insect cultures.  Usually, to keep a temperature this close to ambient room temperature, the equipment needs refrigeration capabilities.

## Culture media

- “Old Standards”
- Grace’s
- Schneider’s
- Mitsuhashi and Maramorosch
- and others

- Additives
- FBS and/or other complex additives
- Growth factors (?)
- Hormones
- Ecdysone
- JH
- Reduced glutathione, cysteine or phenylthiourea
- Nutrients
- Conditioned medium
- Hemolymph
- Antibiotics

- Commercial Serum-free
- Ex-Cell TM 400 series
- Sf-900 II
- Insect-XPRESS TM
- SFX-Insect TM
- Drosophila-SFM
- and others

**Notes:**
Many other formulations exist in addition to the media listed on this slide.  Hink’s TNM-FH is Grace’s modified with the addition of yeastolate, lactalbumin hydrolysate, and fetal bovine serum.  TC-100 (aka BML/TC-10) is also modified from Graces with the reduction/elimination of a couple of amino acids, Krebs cycle intermediates and the addition of tryptose broth.  Goodwin and Vaughn developed a number of media (IPL-41 being the most commonly used) as well. While some of us still use these formulations, especially to maintain cells that were originally established on them, we find the use of some of the serum-free formulations (to which we frequently add serum) consistently effective for primary cultures from a variety of insects.
Some of the additives listed here have rarely been used for developing insect cell cultures (for example, very little is known about growth factors for many specific insect tissues).  In the earliest days of insect cell culture, insect hemolymph was frequently used as a supplement that may have provided some such factors but can also be problematic by introducing detrimental substances.  FBS was an effective substitute but insect-specific factors are likely to be quite different from those from mammals.  In addition to FBS, other ‘undefined’ additives that have been used include protein hydrolysates, serum albumin, and tryptose broth.  Insect hormones, especially ecdysteroids, are growth promoting at low concentrations and cause differentiation at higher (molt-inducing) levels.
Antibiotics are nearly essential for primary culture work but should be discontinued as soon as possible after consistent cell growth is achieved.  

## Source of Cells

- Eggs (embryos)
- Many cell types are actively dividing and undifferentiated
- Whole larvae (neonate)
- All cell types
- Some (most?) are already terminally differentiated
- Larval tissues (from older larvae)
- Specific cell types
- Many terminally differentiated
- Adult tissues
- Reproductive tissues (esp. ovaries)

**Notes:**
See subsequent slides for details of these cell sources.

## Can use various ages of embryos
Clean and disinfect eggs 
	(with 70% ethanol and/or other disinfectants )

- Macerate eggs with ‘mortar and pestle’

**Embryos – method 1**

- Can use various ages of embryos
- Clean and disinfect eggs
- 	(with 70% ethanol and/or other disinfectants )

- Centrifuge to separate tissues from yolk and debris
- Transfer to culture flask
- If culture contains a lot of debris, replace medium at 24 hr.

**Notes:**
Researchers have successfully used variously aged embryos for developing cell lines.  As a rule, Dwight likes to use fairly young embryos while Cindy uses older ones.  
The exact nature of disinfection largely depends on how ‘dirty’ the eggs are.  Dwight generally just  uses 70% ethanol for 5 to 10 minutes followed by several sterile distilled water rinses.  Cindy adds Triton-X100 to the ethanol while Guido uses 1.0% Sodium hypochloride (chlorox)(3 min) followed by 1.0% Quadramine (sterilizing salt)(5 min), then 70% ETOH (5 min) and finally 3 rinses with sterile distilled water.  
If you have a lot of problems with contamination, use a more rigorous method.

## Can use various ages of embryos
Clean and disinfect eggs 
	(with 70% ethanol and/or other disinfectants )

**Embryos – method 2**

- Can use various ages of embryos
- Clean and disinfect eggs
- 	(with 70% ethanol and/or other disinfectants )

- After disinfection, transfer eggs to culture medium
- Cut or tear open chorion
- Separate embryos from yolk material

- Transfer embryos to standing drop of tissue culture medium
- Cut/tear embryos into 3 to 8 pieces

- OR

## Embryonic cell lines

**Notes:**
Since embryos contain all the cell types that will eventually differentiate into larval and adult tissues, cell lines developed from them can contain a wide diversity of cell morphologies.  This will also be addressed in terms of primary cultures in a later slide (#25) but this slide shows cell lines from four insect species: the cabbage looper (Trichoplusia ni) line IPLB-TN-R2; three lines from the tobacco budworm (Heliothis virescens), IPLB-HvE1a, IPLB-HvE6s, and BCIRL-RP-HVE-CLG6; and two each from the tarnished plant bug (Lygus lineolaris); and Mediterranean flour moth (Ephestia kuehniella).

## Disinfect eggs– same procedure  as for embryo cultures
Place in petri dish on dampened filter paper
Wait for hatch
Place 1.0 ml,  0.25% trypsin on Maximov slide
Place 30+ newly hatched larvae in slide
Cut / mince larvae to very fine pieces
Transfer minced larvae to cent. tube / add 4.0 ml additional trypsin
Incubate 37°C / 10 min
Add 1.0 ml FBS to stop action of trypsin
Triturate  vigorously 
Spin / low speed / 5 min
Resuspend pellet / 3.0 ml growth medium + antibiotics

- Place in petri dish on dampened filter paper
- Wait for hatch
- Place 1.0 ml,  0.25% trypsin on Maximov slide
- Place 30+ newly hatched larvae in slide
- Cut / mince larvae to very fine pieces
- Transfer minced larvae to cent. tube / add 4.0 ml additional trypsin
- Incubate 37°C / 10 min
- Add 1.0 ml FBS to stop action of trypsin
- Triturate  vigorously
- Spin / low speed / 5 min
- Resuspend pellet / 3.0 ml growth medium + antibiotics

**Whole neonate larvae – method 1**

**Notes:**
Guido’s method for neonate (newly hatched first instar) larvae.

## Once larvae hatch, they will crawl toward the light into the medium
Use a pipet or glass rod to crush larvae
Transfer medium to flask as primary culture

**Whole neonate larvae – method 2**

- Disinfect eggs– same procedure  as for embryo cultures

- Add 4 ml culture medium to sterile centrifuge tube

- Place eggs near top

- Wrap top of tube in foil

- Once larvae hatch, they will crawl toward the light into the medium
- Use a pipet or glass rod to crush larvae
- Transfer medium to flask as primary culture

- Melanin inhibitor may be necessary
- (Reduced glutathione, cysteine or phenylthiourea)

**Notes:**
This method was described to Dwight by a Japanese researcher at Tsukuba.  None of us have personally used it but seemed like a good ‘trick’ for collecting neonate larvae for primary cultures.

## Older larvae -methods

- (may also need a sodium hypochlorite pretreatment: 1 to 2 minutes with 50% household bleach plus 1% triton X-100 or other detergent)
- Rinse at least twice with sterile distilled water
- Transfer to culture medium

**Older larvae -methods**

**Notes:**
As with the eggs, the method used for disinfection of larvae should be based on how dirty the insects are.  The same agents can be used as discussed in the notes on Slide 5.
Sequential immersion in 20% Septisol (Vetal Laboratories, St. Louis, MO), 0.1% p-hydroxybenzoic acid methyl ester (Sigma-Aldrich, St. Louis, MO) and 0.1% sodium hypochlorite for 2-3 min each followed by distilled water rinses has been used for particularly dirty insects.

## Dissection

**Dissection**

**Notes:**
After disinfection, the insect is transferred to a waxed-bottom Petri dish (which can be sterilized by autoclaving or disinfected with ethanol) containing enough culture medium to cover the insect, pinned to the bottom, dorsal side up, and an incision made along the dorsal midline.  The incision can be kept open with additional insect pins.

## Cell Source –Larval tissues

- Successful for cell lines
- Reproductive
- Ovaries
- Testes
- Hemocytes
- Fat body
- Imaginal discs
- Midguts
- Nerves

- Not previously used for cell lines
- Malphigian tubules
- Tracheoles
- Salivary glands
- Muscles/Aorta
- Endocrine glands

**Notes:**
The cell sources listed under ‘successful’ are based on literature reports and personal information.  The earliest insect cell lines were established from ovaries (although frequently from pupae and adults rather than larvae).

## Reproductive organs

**Ovaries**

**Reproductive ****organs**

**Notes:**
Ovaries: often are difficult to find in younger larvae, but become very prominent during pupal development.  The easiest way to initially locate them in a specific insect is to find them in a larva just prior to the molt to the pupal stage and then perform additional dissections on progressively younger larvae. Cell lines have been established from larvae, pupae and adults.  Sheath cells may be the source of many of these cell lines but the ovarioles may also provide dividing cells.

## Reproductive organs

**Testes**

**Reproductive ****organs**

**Notes:**
Testes: typically are quite easy to find in larvae.   Sheath cells are the likely source of cultured cells.

## Hemolymph

- Melanin inhibitor may be necessary
- (Reduced glutathione, cysteine or phenylthiourea)

**Hemolymph**

**Notes:**
Hemocytes – easy for setting up a culture.  Disinfect the larva, cut a proleg and allow the blood to drip into a dish or flask of medium.  Melanization is a common problem in primary hemocyte cultures but the phenyloxidase activity is typically is reduced during molt.  Use of phenyloxidase inhibitors are also indicated to help overcome the melanization problem.

## Fat body

**Fat body**

**Notes:**
Fat body – equivalent to the mammalian liver and fat.  This tissue does not seem to actively divide in many lepidopteran larvae but when it works, it can work very well.  Fat body cells are a likely source of growth factors so co-culture with other tissues can improve growth of cells in primary cultures.

## Imaginal discs

**Imaginal discs**

**Notes:**
Imaginal discs – aggregates of progenitor cells for adult tissues.  Found in larvae, they are ‘determined’ to become a specific adult structure but are still developmentally undifferentiated.  This makes them very good candidates for cell line development.  Two problems – they are difficult to find and (like hemocytes) may have significant levels of phenyloxidase activity requiring the use of inhibitors in the primary cultures.

## Digestive tract

**Digestive tract**

**Notes:**
Digestive tract – a very important tissue for efforts at insect control.  Often is the first line of defense for pest control, especially pathogens.  The two biggest issues for culture work: sterility and production of digestive enzymes.  Insects in the process of molting usually reduce the amount of gut contents which may aid in obtaining contamination-free cultures.  The use of antibiotics in the insect diet or develop axenic (microbe-free) insect colony may also be effective techniques.  

## Nervous system

**Nervous system**

**Notes:**
Nerves – relatively easy to dissect but typically have few dividing cells.

## Tracheal system

**Tracheal system**

**Notes:**
Tracheoles – easy to dissect.  Should be a useful tool in the study of baculoviruses (one of the earliest tissues to become infected) but probably few dividing cells.

## Salivary glands

**Salivary glands**

**Notes:**
Salivary (silk) glands –easy to find; probably contain few dividing cells.  

## Aorta

**Aorta**

**Notes:**
Muscles - easy to find; probably few dividing cells. 

Quiot et al. [1985. In Vitro Cell. Dev. Biol. 21:603] reported on a dorsal vessel (aorta) cell line from cockroaches but we’ve still included muscles under the ‘non successful’ tissue category since it is such a rare event.  

The cell line pictured here was developed from Trichogramma exiguum embryos and, after the cells had been in culture for some period, was determined capable of differentiating into contractile cells. 

## Endocrine glands

**Endocrine glands**

**Notes:**
Endocrine glands – can be difficult to find; contain few or no dividing cells.  Most work with these tissues have been as primary cultures. 

## Primary Culture Techniques:Tricks of the Trade

- Keep a high “tissue-to-media volume” ratio
- Combine explants from many individuals
- and/or
- Use a standing drop of medium for the first 24-48 hours
- Supply fresh medium on a fairly regular (7-10 day) interval
- Grace’s “organized neglect”

**Notes:**
The ‘feeding’ of primary cultures can be a critical issue in developing cell lines.  
Cindy uses a replacement procedure in which half of the culture medium is replaced at a regular interval.
Dwight generally adds fresh medium to the original cultures over the first few weeks. Using a 35mm Petri dish, the initial standing drop is typically 150-200μl which is increased to 1ml within the first couple of days.  Subsequently, 0.5ml fresh medium is added at 7-10 day intervals until the volume reach ~2.5ml.  At this point, all but 0.5ml of the medium is removed (typically transferred to a culture flask) and 0.5ml fresh medium is added back to the culture.  This addition/replacement routine is continued until the cells can be subcultured.

## Primary Culture Techniques:Tricks of the Trade (Cont.)

- Mechanical vs. enzymatic disruption of tissues
- No single ‘right’ method
- Microscalpel, microscissors or mortar/pestle for mincing/macerating
- or
- Two fine-tipped forceps for tearing
- or
- Trypsin, Collagenase, Hyluronidase, etc. for ezymatic disruption

## Primary Culture Techniques:Tricks of the Trade (Cont.)

- Primary Culture Techniques:Tricks of the Trade (Cont.)

- Selection of colonies based on morphology

- All of these cells were present in a single early passage embryo culture

## Primary Culture Techniques:Tricks of the Trade (Cont.)

- Selection of colonies based on morphology
- Make a cell scraper from a pipet tip

- Use flattened edge to scrape off cells, then suction into tip with pipettor
- Transfer to a new dish or multiwell plate

## Primary Culture Techniques:Tricks of the Trade (Cont.)

- Suspended vs. attached cell selection
- Gentle rinse and transfer for suspended
- Flushing, scraping, and/or enzymes for attached

**Notes:**
This selection method can be for different potential uses (i.e., attached for plaque assays; suspended for large scale).

## Primary Culture Techniques:Tricks of the Trade (Cont.)

- Temperature
- 26-28°C for most temperate insect species
- 16-18° may improve maintenance of preferred traits (virus susceptibility, for example)

**Notes:**
Low temperature lines maintained susceptibility to granuloviruses.

## Once a normal subculture routine is established, leftover cells from each passage can be stored for a few weeks at a lower temperature.   
Add fresh medium to the leftover cells in the parent culture
Leave the parent at room temperature or use a cool incubator (such as the 16-18°C incubator used for low temperature cells).

- Add fresh medium to the leftover cells in the parent culture
- Leave the parent at room temperature or use a cool incubator (such as the 16-18°C incubator used for low temperature cells).

- Backup cultures (short term storage)

## Long term storage

- New cell lines should be stored in liquid nitrogen at the lowest possible passage.
- Record cell identity, passage level, medium, supplements, cryoprotectant used, number of ampoules prepared, name of person doing the freeze, location in the freezer (Freezer no., Canister, Cane) in log book
- Multiple storage locations is recommended

**Notes:**
Freeze procedure:
Cell culture in log-phase growth is suspended into freezing medium (This is typically the culture medium ‘normally’ used for the cells with the addition of 5-10% glycerol or 5-10% DMSO.  If the culture is normally grown in serum-free medium, the addition of serum can also improve recovery.)
1-4 ampoules can be prepared with 1.0ml cell sample from each 25-cm2 flask (Use sealed glass if ampoules are to be submersed in LN2)
Controlled-rate freezing apparatus chills at -1°C/min over critical range
Recovery:
Remove ampoule from freezer while wearing eye protection
Transfer to warm (~30°C) beaker of water
When thawed, dip in 70% ethanol
Dry on clean paper towel
Break ampoule
Transfer contents to centrifuge tube with 10ml fresh medium (same formulation as used prior to freeze)
Centrifuge 50-100Xg, 10 minutes,
Decant medium, resuspend cell pellet into 4ml fresh medium
Transfer to culture flask
Examine after 24 hr, replace medium if much debris
Initial subcultivations should be preformed as necessary