We describe a novel quantitative cDNA expression profiling strategy, involving amplification of the majority of mouse transcriptome using a defined set of 44 heptamer primers. The amplification protocol allows for efficient amplification from as low as 50pg of mRNA and did not alter the expression of the transcripts even with 200 fold dilution of the minimum requirement of the starting material (10ng of mRNA) for standard RNA-seq protocols. We implemented our methodology on embryological lineage segregation, achieved by graded activation of Activin A/TGFß signaling in mouse embryonic stem cells (mESCs). The fold changes in transcript expression were in excellent agreement with quantitative RT-PCR and we observed a dynamic range spanning more than five orders of magnitude in RNA concentration with a reliable estimation of low abundant transcripts. Our transcriptome data identified key lineage markers, while the high sensitivity showed that novel lineage specific transcripts anticipate the differentiation of specific cell types. We compared our strategy with Std. RNA-seq (Mortazavi et al. 2008) and SMART-seq (Ramsköld et al. 2012). We also showed potential of our methodology to suppress the representation of highly expressing ribosomal transcripts. Overall design: Sequencing was performed on day 4 differentiating mouse ESCs treated for two days with 3 different dosages of Activin A (3ng/mL, 15ng/mL and 100ng/mL). The cells were also treated with SB-431542. Serial dilutions of mRNA derived Activin A(3ng/mL) samples were used to detemine the minimum amount of mRNA required to construct relaible sequencing library. SMARTseq libraries were prepared for both Activin A(3ng/mL) and Activin A(100ng/mL) samples. Three Different primer sets were designed to suppress the representaiton of Ribosomal transcripts.