Raw microarray expression data was log2 transformed in SAS.
Arrays were pre-scanned at 0.05% saturation tolerance, and then manually adjusted to minimize dye differences. Images were saved as a .tif file. (Parameters: Scanning hardware = GenePix 4000B [Axon Instruments], Scanning software = GenePix [Axon Instruments])
Oligo Slide Preparation Note: slides can be crosslinked and stored for about a month before pre-hyb and hybridization steps 1. Blow off dust from slide 2. Boil water in a beaker so that a steamy mist is rising from the water. With printed side down (barcode is on printed side), pass the array quickly through the steam and place the slide array side up on a hot block until dry. (Do not hold the slide over the steam for more than a second as the spots will swell and bleed together if they are oversaturated). 3. Place the slides in a UV linker. For the Stratalinker set the exposure to 6000 x 100uJ/cm2. Press "Start" to bind. It should take about 5 minutes to crosslink oligos to the slide. 4. Before prehybridization, plunge slides in 55C 0.2% SDS and immediately shake vigorously for 2 minutes. Shaking prevents excess DNA from adhering to the untreated glass surface, which may cause "comets". Wash twice in pure water about 30 seconds each wash. Transfer to water for 15 sec. and spin dry at 500 x g for 3 minutes. (Spin in a swing bucket rotor. It is important to do this quickly. Bring the third water wash with the slide to the centrifuge to reduce any drying before the spin. Make sure your gloves are dry and clean, any droplets on the side of the slide will spread across the array and leave high background.) Microarray Prehybridization 1. Prepare prehybridization solution. (note: use prehyb twice and then discard) Stock Add Final Concentration 20x SSC 12.5 ml 5x 10% SDS 0.5 ml 0.1% I-block 0.5 g 1% Water 37 ml 2. Prewarm solution at at 42C for ~1 hr to dissolve I-Block. 3. Incubate microarray in prehyb buffer at 42C for 1-1.5 hr. 4. Wash microarray in 3 quick water washes. Use Milli-Q water. 5. Spin microarray in centrifuge to dry (500 g for 3 min). Hybridization 1. Thaw/Resuspend Genisphere formamide hybridization buffer by incubating at 55C for 10 min. 2. Prepare hybridization buffer: Reagent Volume (ul) 2x formamide hyb buffer (Genisphere) 27.5 Combined labeled aRNA 11 KREAblock 13.75 Nuclease-free H2O 2.75 Total Volume 55 3. Incubate at 80C for 10 min. 4. Incubate at 42C until samples are loaded onto microarray 5. Prewarm microarray on heat block at ~40C 6. Hybridize samples (don’t forget to add H2O in the wells of the hyb chamber) 7. Incubate overnight in a 42C waterbath (~15 hr) Post-Hybridization Washes Note: use washes 2x for 6 slides and then replace 1. Prepare hybridization washes and pre-warm wash 1 beaker and tube to 42C. 2. Coverslip removal: beaker of 2x SSC/0.2% SDS at 42C 3. Wash 1: 50ml tube of 2x SSC/0.2% SDS at 42C for 10-12 min 4. Wash 2: 50ml tube of 2x SSC at RT for 10-12 5. Wash 3: 50ml tube of 0.2x SSC at RT for 10-12 6. Centrifuge slides at 500 x g for 2 min. Buffers and Washes All buffers and washes should be filtered before use. Prehybridization Buffer Stock Add Final Concentration 20x SSC 12.5 ml 5x 10% SDS 0.5 ml 0.1% *I-block 0.5 g 1% Water 37 ml *Add the I-block after filtration. Hybridization buffer 50% deionized formamide 5 mL formamide 10x SSC 5 mL 20x SSC 0.2% SDS 200ul 10% SDS Wash 1 100 mL 20x SSC 20 mL 10% SDS 880 mL MilliQ Water Wash 2 100 mL 20x SSC 900 mL MilliQ Water Wash 3 10 mL 20x SSC 990 mL MilliQ Water (Parameters: Chamber type = Corning Microarray Technology- CMT-Hyb chamber, Quantity of label target used = 5, Mass unit = Nano gram, time = 15, Tiny time unit = hours, Volume = 55, Volume unit = Micro litre, temperature = 42)
Labeling and Fragmentation MATERIALS Kreatech ULS-Cy 3/5 aRNA Labeling Kit (purchased from Open Biosystems – KRE4016). ULS-Cy 3/5 aRNA Labeling Kit – Open Biosystems #KRE4016 Ambion Fragmentation Reagents – Ambion #8740 ULS Labeling 1. Use 4 ug of antisense, amplified RNA in each reaction with a total concentration > 50ng/ul. 2. For each reaction, place the following reagents in an RNase-free microcentrifuge tube: Reagent Volume (ul) aRNA (4 ug) ? 32 Cy3/5-ULS 4 10x labeling solution 4 Nuclease-free H2O up to 32 Total Volume 40 3. Mix briefly and centrifuge to collect mixture at the bottom of the tube. 4. Incubate at 85C for 30 min. 5. Place samples on ice. 6. Centrifuge briefly and place back on ice. Labeled aRNA Purification (KREApure columns) 1. Resuspend KREApure column material by vortexing. 2. Loosen cap ¼ turn and snap off bottom closure. 3. Place column in 2 ml collection tube. 4. Pre-spin column for 1.5 min at 20,800 x g. 5. Discard flow through 6. Wash column with 300 ul RNase-free H2O. 7. Spin 1.5 min at max speed, discard flow through. 8. Spin 1.5 min at max speed, discard flow through and collection tube. 9. Transfer column to new microcentrifuge tube. 10. Add ULS-labeled sample to middle of column bed. 11. Spin column for 1.5 min at 20,800 x g. 12. The elutate contains the purified labeled aRNA (~38 ul). Analysis of Labeled aRNA 1. Use Nanodrop to spec 1 ul of sample using microarray settings. 2. Calculate volume for each sample to contain 120 pmol dye. 3. Pool 120 pmol dye from each paired Cy3/Cy5 sample and put into 2.0 ml tube. 4. Dry pooled samples down to 9 ul. 5. Transfer to 0.2 mL tube aRNA Fragmentation (Ambion Fragmentation Reagents) 1. Add 1 ul of 10x fragmentation buffer to 9 ul of sample (1/10 volume fragmentation buffer). 2. Incubate at 70C for 15 min. 3. Briefly spin tube. 4. Add 1 ul stop solution and mix by pipetting. 5. Place on ice or freeze at -80C until hybridization (Parameters: Amount of nucleic acid labeled = 4, Mass unit = Micro gram, Amplification = none)
Required kits: RNeasy RNA extraction kit (Qiagen, Valencia, CA), RNase-free DNase 1. Protocol: 1. Dissect bee abdomens on ice and in RNAlater and remove any internal organs. Use RNAlater for preserving RNA (200ul). At this point cuticles with attached fat bodies can be stored at -80 degrees C. 2. Put frozen samples on (regular) ice to thaw. Prepare RLT/beta-mercaptoethanol solution (solution 1) (1ml of RLT and 10 ul of beta-ME). Need 600 ul of solution 1 per sample. 3. Remove 200 ul of RNAlater from the sample, add ONLY 200ul of solution 1 to the sample. Homogenize carefully with the mortar and pestle. At the end clean homogenizer with 400 ul of solution 1. Put the samples on dry ice. At this point samples can be stored at -80 degrees C. 4. Centrifuge lysate for 3 min. at max speed (13200rpm) in a microcentrifuge. 5. Transfer supernatant to a new tube (there should be ~600 ul for abdomen-fat body). Use in subsequent steps. 6. Add 600 ul of 70% ethanol to the supernatant and mix by pipetting . DO NOT CENTRIFUGE! 7. Apply up to 600 ul (so 2X for us) of the sample (including any precipitate formed) to an RNeasy mini column placed in a 2 ml collection tube (supplied). Close the tube and centrifuge for 15 s at 10000rpm. Discard the flow-through and reuse the collection tube in the next step. 8. Pipet 350 ul RW1 buffer into an RNeasy column (directly onto a filter) and centrifuge 15 s at 10000rpm. Discard flow-through. 9. Add 10 ul DNase I stock solution to 70 ul RDD buffer. Mix by gently inverting the tube. DO NOT VORTEX! Note: To make DNase I stock solution dissolve the solid DNase in 550 ul of RNase free water. Mix gently. DO NOT VORTEX! 10. Pipet the DNase I/RDD mix (80 ul) directly onto the RNeasy silica-gel membrane and leave for 15 min. at room temperature. 11. Pipet 350 ul RW1 buffer into RNeasy mini column. Centrifuge for 15 s at 10000rpm. Discard flow-through. 12. Transfer the RNeasy column into a new 2ml tube (supplied). Pipet 500 ul RPE buffer onto RNeasy column. Close the tube gently and centrifuge for 15 s at 10000rpm. Do not shake! Discard the flow-through. 13. Add another 500 ul RPE buffer to the RNeasy column. Close the tube gently and centrifuge for 2 min at 10000rpm to dry the membrane. 14. Place the RNeasy mini column into a new 2 ml tube (not supplied) and discard the old tube. Centrifuge for 1 min at full speed. 15. To elute, transfer the RNeasy column into a new 1.5 ml collection tube (supplied). WAIT 1 MIN!! Pipet 40 ul RNase free water directly onto silica-gel membrane. Close the tube gently and centrifuge for 1 min at 10000rpm. Note: This step may be repeated if RNA concentration is not high enough. Elute into same collection tube. 16. Measure the amount of RNA with a NANO DROP (2 ul of sample). 17. Keep the eluted RNA stored at -80 degrees C.
Honey bee queens (Apis mellifera carnica) used were reared at the Honey Bee Research Facility, North Carolina State University (Raleigh, NC). The source colony used for queen grafting was headed by a single-drone inseminated queen (Glenn Apiaries, Fallbrook, CA). Since honey bees exhibit haplodiploidy, the queens used were highly related with the average coefficient of relatedness (G) of 0.75. Young worker larvae were grafted into commercially available queen cups (Brushy Mountain Bee Farm, Moravian Falls, NC) with royal jelly and placed in queenless colonies to be reared according to standard practices (Laidlaw, 1977; Laidlaw and Page, 1997). A week after grafting, sealed queen cells were placed in an incubator (33C and ~50% RH) where they remained for 48 hours. Two days prior to their expected emergence, queen cells were placed into individual mating nuclei (Brushy Mountain Bee Farm, Moravian Falls, NC) with approximately 1,000 workers, one frame of young brood, one frame of honey and pollen, and one empty frame. Seven days post-emergence, all of the queens were marked with a marking pen (Dadant and Sons, Inc., Hamilton, IL) and a number tag (Betterbee, Inc., Greenwich, NY) on their thorax, their left wings were clipped, and they were placed in one of five randomly assigned groups: untreated virgin control (8 queens), instrumentally inseminated with either 1ul (9 queens) or 8ul (11 queens) of saline (referred to as Sa1 and Sa8 groups respectively), and either 1ul (11 queens) or 8ul (9 queens) of semen (referred to as Se1 and Se8 groups respectively). (Parameters: time unit = seconds, temperature unit = C)
Virgin queens did not receive any treatment, but they were handled, marked, and clipped on the same day that the other queens were instrumentally inseminated. Saline solution was prepared as in (Williams Harbo 1982)and used to inseminate the queens in Sa1 and Sa8 groups. A mixed pool of semen was prepared from approximately 150 drones collected at hive entrances upon their return from failed mating flights and stored overnight in cages placed inside the hives so the workers still had access to feed the drones. Semen was collected in the morning on the day of instrumental inseminations. Semen was gently mechanically mixed and used for all inseminations of queens in the Se1 and Se8 groups. After manipulation and/or insemination, queens were returned to their respective mating nuclei. Five queens were used for microarray analysis from each group and only the queens with highly activated ovaries (ovary scores of 3 or 4) were used for the treatment groups. Virgin queens had ovaries exhibiting no or slight activation (ovary scores of 1 or 2).